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BIOMERIT Research Centre, Department of Microbiology, University College Cork, Ireland
Correspondence
Fergal O'Gara
f.ogara{at}ucc.ie
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). Given that epithelial cells act as the initial barrier to infection, an understanding of the interactions between these cells and the bacterium is critical. P. aeruginosa has also been shown to cause infections in non-mammalian hosts such as insects (Jander et al., 2000) and plants (Rahme et al., 1995; Silo-Suh et al., 2002; Walker et al., 2004). The metabolic versatility of this bacterium is reflected in the fact that 10 % of its genome encodes membrane-associated transporters, with a further 8 % involved in the metabolism of carbon-containing compounds (Stover et al., 2000). One transport system of particular interest is the dipeptide (dpp) transport operon because of its involvement in the transport of not only dipeptide containing compounds but also aminolaevulinic acid (Carter et al., 2002; King & O'Brian, 1997), haem (Letoffe et al., 2006) and single amino acids (Tamber & Hancock, 2006).
The dpp transport operon has been well characterized and shows conservation in structure across both Gram-negative (Abouhamad & Manson, 1994) and Gram-positive (Guedon et al., 2001; Slack et al., 1995) bacterial species. The dpp operon has been shown to be maximally expressed in the stationary phase of growth, with expression driven from the promoter region upstream of the first gene, dppA (Abouhamad & Manson, 1994). In Salmonella typhimurium and Escherichia coli, dppA has been shown to play a role in chemotaxis (Abouhamad et al., 1991). The role of the dpp transport machinery in utilizing haem iron has also been shown in E. coli. Binding of haem by the DppA protein allows transport of this molecule into the cell via the dppBCDF ATP-binding cassette transporter (Letoffe et al., 2006).
To date relatively little information is available regarding the regulation of this transport machinery. Regulation of the dpp transport operon in Bacillus subtilis has been shown to be exerted by the pleiotropic negative regulators CodY and AbrB (Slack et al., 1995). In Lactococcus lactis, a homologue of CodY was shown to respond to the levels of branched-chain amino acids, thereby sensing aspects of the nutritional supply for the cell. Additionally an lrp (encoding a leucine-responsive regulatory protein) gene from Bradyrhizobium japonicum has been shown to complement an E. coli dpp mutant strain for the uptake of dipeptides (King & O'Brian, 1997). In E. coli the gcvB gene, which encodes a small untranslated RNA, has been shown to regulate expression of both the Dpp and oligopeptide (Opp) transport systems. This small untranslated RNA appears to regulate genes associated with oligopeptide transport at the translational level whereas it appears to regulate dppA expression at the mRNA level (Urbanowski et al., 2000).
P. aeruginosa PAO1 contains a gene cluster which shows a high level of homology to previously characterized dpp transport operons (http://www.pseudomonas.com). Interestingly, transcriptome analysis has shown that dppA3 in P. aeruginosa PAO1 is induced under low-iron conditions (Ochsner et al., 2002). The porin OpdP, which is associated with this putative dipeptide transport machinery in P. aeruginosa PAO1, is required for the transport of the dipeptide glycyl-glutamic acid (Tamber & Hancock, 2006). The opdP gene is induced to a high level when bacteria are grown on the amino acid L-arginine, suggesting involvement of OpdP in the uptake of single amino acids as well as dipeptides (Tamber & Hancock, 2006).
Previous transcriptome analysis of P. aeruginosa, carried out in the presence of a human airway epithelial cell line, showed that expression of the gene PA4498, encoding a putative metallopeptidase, was upregulated 4.8-fold (Frisk et al., 2004). Variations in gene expression have been shown to influence the virulence of the Liverpool epidemic strain (LES) of P. aeruginosa. The highly virulent, epidemic strain LES431 has shown increased resistance to antimicrobials and has caused mortality in both immunocompromised and non-immunocompromised patients. Transcriptome analysis of this strain has shown that it also overexpresses the mdpA gene, encoding a putative metallopeptidase (Salunkhe et al., 2005).
This paper identifies the psdR gene as a transcriptional repressor of the mdpA and dppA3 genes. It also demonstrates that the mdpA gene is essential for the growth of P. aeruginosa on dipeptides as sole carbon source and suggests an involvement of this gene in P. aeruginosa cytotoxicity.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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. All cultures of P. aeruginosa PAO1 and associated mutants were routinely grown on Casamino Acid (CAA) medium supplemented with 100 µM FeCl3, LB or M9 minimal medium (Sambrook et al., 1989) at 37 °C. The mPA4498 (mPAmdpA) mutant strain was obtained from the University of Washington Mutant Library (Jacobs et al., 2003). E. coli strains were routinely grown in LB medium at 37 °C. Where appropriate, antibiotics were added to growth media at the following concentrations: for P. aeruginosa, 50 µg kanamycin ml–1, 200 µg chloramphenicol ml––1 and 50 µg gentamicin ml–1; for E. coli, 25 µg gentamicin ml–1, 50 µg ampicillin ml–1 and 12.5 µg tetracycline ml–1.
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; Rajan et al., 2000). Cells were cultured on BSA-collagen-fibronectin-coated plastic, using minimum essential medium supplemented with 10 % fetal bovine serum, 2 mM L-glutamine, 100 µg penicillin ml–1, 100 µg streptomycin ml–1 and 400 µg neomycin G-418 ml–1. Cells were incubated at 37 °C in a humidified 5 % CO2 atmosphere. All cell culture reagents were supplied by Sigma, unless stated otherwise.
DNA and RNA manipulations.
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 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 according to the manufacturer's specifications (Invitrogen). Plasmids were mobilized into Pseudomonas strains by triparental mating using the helper plasmid pRK2013 (Figurski & Helinski, 1979).
Determination of nucleotide sequence and sequence analysis.
All transcriptional fusions and psdR gene disruption were confirmed by nucleotide sequencing performed at 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 Centre for Biotechnology Information (NCBI). Multiple sequence alignments were performed using the CLUSTAL W program (Chenna et al., 2003).
Construction of the psdR mutant.
To construct a PA4499 (psdR)-negative mutant we first amplified two genomic fragments. Fragment 1 (Frg1) covers the region extending from +1435 to –130 relative to the PA4498 (mpdA) ATG start codon, and fragment 2 (Frg2) covers the region extending from +270 relative to the psdR ATG start codon to +507 relative to the PA4500 (dppA3) ATG start codon. These two fragments are located 348 bp apart. The joining of Frg1 and Fgr2 would delete 348 bp of the psdR gene including 58 bp of the untranslated region upstream to the ATG and the first 96 codons of the psdR ORF. We engineered PstI and NotI restriction sites at the 5' and the 3' ends of Frg1 and NotI and XbaI restriction sites at the 5' and the 3' ends of Frg2 respectively. These fragments were separately cloned into the TOPO cloning vector. Frg1 and Fgr2 were then joined at the NotI site. The vector containing Frg1 and Frg2 was linearized at the NotI site and ligated to the gentamicin-resistance cassette digested with the same restriction enzyme to produce the recombinant plasmid P
psdR1 containing a disrupted psdR gene. The disrupted gene was transferred to into the allelic-exchange vector pK19mobsac using PstI and XbaI restriction sites to create pK19psdR. The sacB gene contained on pK19mobsac codes for the enzyme levansucrase, which confers sucrose susceptibility to cells expressing the intact gene. Triparental mating was carried out to transfer this construct from E. coli XL1 to P. aeruginosa PAO1 wild-type strain. The positive clones, which had integrated the plasmid into the chromosome, were selected for on 50 µg gentamicin ml–1, 50 µg ampicillin ml–1. The loss of the sacB gene by a second crossover allows the host to grow on medium supplemented with 5 % sucrose. Thus to select for a double crossover event we plated all PAO1 transconjugants that were gentamicin resistant on LB supplemented with 5 % sucrose. All gentamicin- and sucrose-resistant colonies were considered as candidates to possibly contain the double crossover. Cells were subsequently screened by PCR using primers PK4499CKF and PK4499CKR (Table 2). The confirmed disruptants with the double crossover were named PApsdR.
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Construction of the transcriptional fusions.
To characterize the transcriptional activity of the psdR, mdpA and dppA3 genes, we used the pMP220 vector bearing a promoterless lacZ reporter gene. Two pairs of primers, PKTF4499F/PKTF4499R and PKTF4498F/PKTF4498R (Table 2
), were used to amplify a 592 bp fragment containing the psdR–mdpA intergenic region with the psdR and mdpA predicted promoters and part of the ORF of both genes. To facilitate directional cloning into pMP220 and a correct fusion of psdR and mdpA promoters with the lacZ reporter gene we introduced a PstI site downstream and an EcoRI site upstream of each promoter. Therefore, both transcriptional fusions, pTF99 (psdR) and pTF98 (mpdA), contain the full-length mdpA–psdR intergenic region but in different orientation. The two constructs were transferred into P. aeruginosa PAO1 and the PApsdR mutant strain by electroporation to produce pMP220 : : PpsdR and pMP220 : : PmdpA respectively. To characterize the transcriptional activity of dppA the primers PKTF4500F and PKTF4500R were used to amplify a 414 bp region. This consisted of the entire psdR–dppA intergenic region and the first 62 bp of the dppA gene. To facilitate directional cloning into pMP220 and correct fusion of the dppA promoter with the lacZ reporter gene, a downstream PstI and upstream EcoRI site were introduced. This construct was then transferred into P. aeruginosa PAO1 and the PApsdR mutant strain by electroporation to produce PAO1/pMP220 : : PdppA and PApsdR/pMP220 : : PdppA respectively.
To verify the transcriptional fusion analysis the strains PApsdR/pMP220 : : PdppA and PApsdR/pMP220 : : PmdpA were complemented using the vector pBBRpsdR. The construct was transferred into PApsdR/pMP220 : : PdppA and PApsdR/pMP220 : : PmdpA by electroporation to produce PApsdR/pMP220 : : PdppA pBBRpsdR and PApsdR/pMP220 : : PmdpA pBBRpsdR respectively.
Proteomic analysis
Protein isolation.
Cells were inoculated at an OD600 of 0.01 and grown to an OD600 of 0.6 in 200 ml CAA medium. Total protein from the bacterial suspension was isolated by removing whole cells from the culture by centrifugation (7000 r.p.m., 10 min at 4 °C) using a Sorvall SS-34 rotor in a Sorvall RC-5B centrifuge. The pellet was resuspended in 6 ml HEPES buffer (0.1 M, pH 7.4), protease inhibitor cocktail, DNase (50 U), RNase (10 mg) and 200 µl MgCl2 (100 mM). Resuspended cells were lysed by sonication (5x20 s). Cellular debris was removed by centrifugation (6000 r.p.m., 10 min at 4 °C). Aliquots of 1–2 ml of extracts were extracted twice with 1 ml phenol heated for 10 min at 70 °C. The proteins (solid phase) were precipitated by adding 2 ml acetone.
Two-dimensional gel electrophoresis.
Pellets were resolubilized in solution A [7 M urea, 2 M thiourea, 4 % (w/v) CHAPS, and 50 mM DTT] and diluted to a concentration of 10 mg protein ml–1. Then 100 µl solution B [7 M urea, 2 M thiourea, 4 % (w/v) CHAPS, 50 mM DTT, carrier ampholytes, Triton X-100 and bromophenol blue (0.005 g ml–1)] were added to 3 mg protein. Proteins were loaded by in-gel rehydration on 24 cm pH 4–7 immobilized pH gradient (IPG) strips (Amersham Biosciences). Isoelectric focusing was conducted for a total of 89 kV h. IPG strips were then reduced, alkylated and loaded for second-dimension SDS-PAGE as described previously (Nouwens et al., 2000). Coomassie-blue-stained gels were imaged using a Fluor-S Multimager. To reduce the likelihood of false variations in spot intensity, protein mix from each sample was separated in triplicate gels. The 2D gel electrophoresis was repeated twice in two different experiments. To reduce gel-to-gel variation within the three repeated gels for each strain, one average gel for the proteome of each strain was generated using Photoretix PG200 software (Nonlinear Dynamics).
Peptide mass mapping.
Protein spots were excised from the gel, washed three times with 120 µl ammonium bicarbonate (pH 7.8, 25 mM)/acetonitrile 50 % (v/v) and dried using a SpeedVac (Savant Instruments). Gel pieces were rehydrated with 8 µl ammonium bicarbonate (25 mM, pH 7.8) containing 10 ng porcine modified trypsin µl–1 (Promega) for 10 min at room temperature. Proteins were then digested at 37 °C for 16 h. Peptides were extracted from the gel in 8 µl 50 % acetonitrile/1 % trifluoroacetic acid (TFA). A 1 µl aliquot of each extract was spotted onto a target plate, covered with 1 µl 8 mg 
-cyano-4-hydroxycinnamic acid ml–1 in 50 % acetonitrile/1 % TFA and allowed to dry. Peptide masses were generated by matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF MS) using a Maldi-TOF Voyager DE-Pro. All spectra were obtained in reflectron mode using an accelerating voltage of 20 kV. Mass calibration was performed using Calibration Mix 2 (PE Biosystems) as an external standard. The peptide mass profile was analysed using Data Explore (PerSeptive Biosystems) software and tested against the Mascot Peptide Mass Fingerprint database (http://www.matrixscience.com). The parameters for a successful identification included a mass tolerance of 150 p.p.m. per peptide and total sequence coverage greater than 30 %. Identified proteins were reconfirmed by checking their predicted mass and pI values against their location in the gels.
Epithelial cell infection conditions.
For infection studies, cells were washed twice with phosphate-buffered saline, detached and resuspended in the described cell culture medium and seeded at the appropriate cell number 5 days prior to infection to obtain confluent monolayers on the day of infection. Bacterial strains were cultured overnight aerobically for 16–18 h in cell culture medium without antibiotics (infection medium) at 37 °C. Following PBS washing to remove extracellular components, bacterial densities (in infection medium) were adjusted so as to infect PBS-washed, fully confluent cell monolayers at an m.o.i. of approximately 50 : 1. Serial dilutions were plated onto LB agar to confirm both the m.o.i. used and the bacterial numbers after 9 h of infection. C.f.u. ml–1 values for mutant and wild-type strains showed no significant difference. Quantification of cytotoxicity was carried out for mPAO1 and the mdpA mutant strain as previously described (O'Grady et al., 2006).
Growth curve analysis.
Mutants and parent strains were grown overnight in M9 minimal medium (Sambrook et al., 1989) supplemented with glucose. Cells were then washed in quarter-strength Ringer's solution and resuspended at a final OD600 of 0.01 in M9 minimal medium supplemented with different dipeptides (Gly-Asp, Gly-Gln, Gly-Gly, Gly-Phe, Gly-Ala, Ala-Val, Val-Val, Phe-Pro, Gly-Pro) at a concentration of 10 mM. OD600 readings were taken over a 4 day period (30 s every 15 min) with cultures grown with shaking at 37 °C. Analysis of growth was carried out using the Bioscreen C (Oy Growth Curves, Helsinki, Finland) 200-well culture growth monitoring instrument.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). In particular the cluster of genes associated with dpp transport in P. aeruginosa strains PAO1 and PA14 contains an extra four genes which encode a putative transcriptional regulator (PsdR), a putative metallopeptidase (MdpA) and two predicted dipeptide-binding proteins (DppA1, DppA2). The mdpA and psdR genes are not found in the genomes of Pseudomonas entomophila L48 (Vodovar et al., 2006), P. putida KT2440 (Nelson et al., 2002), P. syringae pv. tomato DC3000 (Buell et al., 2003), P. syringae pv. syringae B728a (Feil et al., 2005) and P. syringae pv. phaseolicola 1448A (Joardar et al., 2005). Bioinformatic analysis of mdpA, located upstream of the putative transcriptional regulator psdR, has identified the protein as a putative metallopeptidase. BLASTP pairwise comparisons showed that MdpA has substantial amino sequence similarity to and domain conservation with Xaa-Pro aminopeptidases. Domain analysis showed that PsdR is a MerR family-like regulator with a helix–turn–helix (HTH) XRE (xenobiotic response element) domain and a cupin sensor domain. In P. fluorescens Pf-5 bioinformatic analysis identified a region with DNA sequence homology to psdR. However, this DNA sequence was found to contain an internal stop codon.
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). In order to reduce gel-to-gel variation within the three repeated gels for each strain, one average gel was generated for each set using Photoretix PG200 software. The use of this approach to compare proteome profiles from wild-type and the psdR mutant stain identified one genuine difference (Fig. 2a, b). The protein spot that was present in the psdR mutant and undetectable in the parent strain was characterized using MALDI-TOF MS and identified as the putative metallopeptidase, MdpA. These results suggest that the putative transcriptional regulator, PsdR, negatively regulates expression of the metallopeptidase, MdpA.
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), to which PsdR shows homology. This observation suggests that the expression of mdpA could be under the control of PsdR. Inspection of the promoter region of dppA3 identified a promoter region homologous to that of mdpA (Table 3). This suggests that PsdR may also function in regulating expression of the dpp transport machinery.
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). This result indicates that expression of mdpA is negatively regulated by PsdR. Expression of dppA3 was significantly higher in the mutant backround, and levels of expression increased throughout growth. This result suggested that dppA3 expression is also negatively regulated by psdR. When the psdR mutant was complemented with the plasmid pBBRpsdR, the level of expression of mdpA was similar to wild-type and remained below 400 Miller units over the 24 h period. A similar pattern emerged when expression of dppA was analysed in a psdR complemented strain, with levels of expression similar to those of the wild-type (data not shown). This verifies that a mutation in psdR results in increased expression of both mdpA and psdR.
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) in P. aeruginosa PAO1 prompted us to hypothesize that MdpA may be involved in the metabolism of this dipeptide. This was examined by assessing the wild-type and an mdpA mutant strain for growth in the presence of different dipeptide carbon sources including glycyl-L-glutamic acid. Wild-type and mdpA mutant strains showed indistinguishable growth in M9 minimal medium supplemented with glucose as a carbon source (data not shown). Unlike the wild-type the mdpA mutant strain was unable to utilize a range of dipeptides as a sole carbon source (Table 4). A psdR mutant strain was also compared to wild-type for growth when these dipeptides were used as sole carbon sources. Disruption of psdR resulted in a shorter lag phase of growth when these dipeptides, including Gly-Glu (Fig. 4b), were used. When X-Pro dipeptides, including Phe-Pro and Gly-Pro (Fig. 4c), were used as carbon sources, the growth kinetics of wild-type and mutant strains were more similar.
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Expression of mdpA was significantly induced, about fourfold, only in the presence of the X-Pro dipeptides (Phe-Pro, Gly-Pro) (Fig. 5). This result indicated that the mdpA gene promoter is directly or indirectly induced by X-Pro dipeptides.
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; Salunkhe et al., 2005). This prompted us to test the role of mdpA in virulence using a cytotoxicity assay on human epithelial cells. Cytotoxicity of airway epithelial cells was measured by quantifying the release of lactate dehydrogenase into culture supernatants. At an m.o.i. of 50 : 1, PAO1 was about four times more cytotoxic to airway epithelial cells than the mdpA mutant at 6 h post-infection (Fig. 6). Thus mdpA appears to contribute to cytotoxicity in PAO1.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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The results reported here show that expression of psdR in the wild-type strain does not fully repress mdpA expression and that the transcription of this gene is growth phase dependent. This suggests that mdpA is under control of other regulators which are not affected by PsdR. Previous experimental evidence suggests that the regulation of expression of mdpA is complex and is controlled by many regulatory inputs. Transcriptome analysis suggests that mdpA is regulated by quorum-sensing-dependent regulators LasR and RhlR in P. aeruginosa PAO1. Expression was shown to be ninefold higher when the PAO1 and lasR/rhlR double mutant strains were compared (Schuster et al., 2003). A similar phenotype is seen in the post-transcriptional regulator rsmA mutant background (Burrowes et al., 2006).
Transcriptome-profiling experiments had previously shown that mdpA is induced both in the presence of an epithelial cell line (Frisk et al., 2004) and in a highly virulent LES isolate of P. aeruginosa PAO1 (Salunkhe et al., 2005). This result suggested to us that this gene could play a specific role in interactions with epithelial cells. Interestingly, a mutation in mdpA results in reduced cytotoxicity of the bacterium to an epithelial cell line (Fig. 6). P. aeruginosa produces several proteases that are implicated in the process of infections caused by this organism (Doring et al., 1985). Most clinical strains of P. aeruginosa produce elastase, a zinc metalloprotease that is implicated in the pathogenesis of infections related to these organisms. Indeed, numerous other examples of proteases and aminopeptidases that are linked to virulence have been highlighted (Coffey et al., 2000; Corbett et al., 2003; Feil et al., 2005; Hidalgo-Grass et al., 2006; Kooi et al., 2006; Song et al., 2004; Spratt et al., 1995). Dipeptidyl aminopeptidase activity has previously been implicated in the virulence of Porphyromonas gingivalis (Kumagai et al., 2000). This aminopeptidase has been shown to function extracellularly. However, sequence analysis of MdpA does not show the presence of a signal peptide involved in triggering protein translocation; it rather suggests a cytoplasmic location for this protein. The exact mechanism of how the dipeptidase MdpA is involved in virulence is unknown.
Analysis of Pseudomonas genomes has shown the presence of the mdpA dipeptidase gene specifically in P. aeruginosa strains and not in other sequenced Pseudomonas species. Questions remain regarding what biological roles both the regulator and peptidase play in Pseudomonas–host interactions. The contribution of MdpA is potentially explained by its role in cytotoxicity, presumably because of the advantages in nutrient metabolism it affords. The physiological role of PsdR, which functions as a repressor of both dipeptide transport and metabolism, is less clear. Perhaps a tighter regulation of genes involved in the uptake and metabolism of small molecules into the cell is required for the successful colonization of environments in which P. aeruginosa is prominent.
It is assumed that the genetic complexity of P. aeruginosa allows for its ecological and metabolic versatility. Analysis of the dpp gene cluster in P. aeruginosa, the characterization of novel genes associated with both nutrient transport and metabolism and the role of mdpA in cytotoxicity would support this view.
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ACKNOWLEDGEMENTS |
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Edited by: M. A. Curtis
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Received 13 November 2007;
revised 2 April 2008;
accepted 11 April 2008.
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