Skip to main page content

• HOME
• Current Issue
• Archive
• Alerts
• About ASM
• Contact us
• Tech Support
• Journals.ASM.org

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Wilhelm, S. Articles by Jaeger, K.-E. Search for Related Content PubMed PubMed Citation Articles by Wilhelm, S. Articles by Jaeger, K.-E.

 Previous Article  |  Next Article 

Journal of Bacteriology, September 2007, p. 6695-6703, Vol. 189, No. 18
0021-9193/07/$08.00+0     doi:10.1128/JB.00023-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

The Autotransporter Esterase EstA of Pseudomonas aeruginosa Is Required for Rhamnolipid Production, Cell Motility, and Biofilm Formation

Susanne Wilhelm,¹ Aneta Gdynia,¹ Petra Tielen,^2, Frank Rosenau,^1* and Karl-Erich Jaeger¹

Institute For Molecular Enzyme Technology, Heinrich Heine University Duesseldorf, Research Centre Juelich, D-52426 Juelich, Germany,¹ Biofilm Centre, University Duisburg-Essen, Geibelstr. 41, D-47057 Duisburg, Germany²

Received 5 January 2007/ Accepted 2 July 2007

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Pseudomonas aeruginosa PAO1 produces the biodetergent rhamnolipid and secretes it into the extracellular environment. The role of rhamnolipids in the life cycle and pathogenicity of P. aeruginosa has not been completely understood, but they are known to affect outer membrane composition, cell motility, and biofilm formation. This report is focused on the influence of the outer membrane-bound esterase EstA of P. aeruginosa PAO1 on rhamnolipid production. EstA is an autotransporter protein which exposes its catalytically active esterase domain on the cell surface. Here we report that the overexpression of EstA in the wild-type background of P. aeruginosa PAO1 results in an increased production of rhamnolipids whereas an estA deletion mutant produced only marginal amounts of rhamnolipids. Also the known rhamnolipid-dependent cellular motility and biofilm formation were affected. Although only a dependence of swarming motility on rhamnolipids has been known so far, the other kinds of motility displayed by P. aeruginosa PAO1, swimming and twitching, were also affected by an estA mutation. In order to demonstrate that EstA enzyme activity is responsible for these effects, inactive variant EstA* was constructed by replacement of the active serine by alanine. None of the mutant phenotypes could be complemented by expression of EstA*, demonstrating that the phenotypes affected by the estA mutation depend on the enzymatically active protein.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Pseudomonas aeruginosa is an increasingly prevalent opportunistic gram-negative pathogen, which can infect humans, but also plants, nematodes, insects, amoebae, and animals (14, 45, 47). In humans, it causes infections in immunocompromised hosts such as patients suffering from cystic fibrosis, burns, or cancer and represents a major problem in intensive care units. P. aeruginosa produces and secretes an arsenal of virulence factors, which are all involved in the initiation or establishment of infection processes. These extracellular virulence determinants include not only proteases, lipases, and phospholipases but also nonprotein compounds like iron scavenger siderophores, the polysaccharide alginate, and the biosurfactant rhamnolipid, which is also known as heat-stable hemolysin (46).

Many of these factors appear to have functions in complex physiological processes related to virulence like adhesion to different surfaces, cellular surface motility, and biofilm formation. P. aeruginosa is capable of performing three different types of cell motility: flagellum-mediated "swimming," type IV pilus-dependent "twitching," and a complex coordinated multicellular migration called "swarming" (16, 39, 48, 54). In contrast to that of many other bacteria, swarming of P. aeruginosa is not dependent only on flagella but also on type IV pili (32). Furthermore, rhamnolipids seem to be required for swarming motility by acting as a surface-modifying agent (32).

The rhamnolipids produced by P. aeruginosa are composed of mono- or dirhamnose linked to 3-hydroxy fatty acids of various chain lengths. The most abundant rhamnolipid species of P. aeruginosa are L-rhamnosyl-3-hydroxydecanoyl-3-hydroxydecanoate (monorhamnolipid) and L-rhamnosyl-L-rhamnosyl-3-hydroxydecanoyl-3-hydroxydecanoate (dirhamnolipid) (37, 50). The rhamnolipid biosynthesis pathway (56) includes two sequential rhamnosyl transfer reactions. The membrane-bound rhamnosyltransferase RhlB catalyzes the formation of monorhamnolipid from the precursors dTDP-L-rhamnose and 3-hydroxyalcanoyl-3-hydroxyalcanoate (HAA). Subsequently, dirhamnolipid can be formed by the RhlC-catalyzed addition of another molecule of dTDP-L-rhamnose to monorhamnolipid (37, 43, 46). The rhlB gene, encoding rhamnosyltransferase 1, is arranged in a bicistronic operon together with rhlA, which encodes an enzyme involved in the synthesis of the fatty acid dimer moiety of rhamnolipids and HAAs (16). HAAs have been identified as precursors in rhamnolipid biosynthesis and are also detectable in P. aeruginosa culture supernatants, possess potent surface-active properties, and affect swarming motility (16, 35).

The biosynthesis and biochemistry of rhamnolipids have been extensively studied; however, the exact function of rhamnolipids is still unclear. They seem to play multiple roles, as their presence promotes uptake of hydrophobic substrates (1, 3, 42) and alters cell surface polarity (1, 11, 66). Rhamnolipids also have antimicrobial activities against other bacteria (19) and are able to disrupt Bordetella bronchiseptica biofilms (25). In P. aeruginosa biofilms rhamnolipids are suspected of playing a role in maintaining fluid channels and the detachment of cells from the biofilm community (7, 15, 18). Furthermore, they appear to act as virulence factors because they severely affect normal tracheal ciliary function (49), inhibit the phagocytic response of macrophages (41), and act as heat-stable hemolysins (26). Swarming motility of P. aeruginosa is affected by rhamnolipids and HAAs, most probably by a reduction of surface tension, which causes the surface conditioning needed for efficient colonization (16, 38).

Several lipolytic enzymes are secreted by P. aeruginosa, including the esterase EstA, which is an autotransporter protein located in the outer membrane (63). Autotransporters predominantly show physiological functions related to the virulence of the corresponding organisms (20). EstA was found to be required for full virulence in a rat model of chronic respiratory infection (45), but its exact physiological function remains to be elucidated.

Since we considered rhamnolipid as a plausible candidate substrate for EstA, we have compared several rhamnolipid-related physiological functions of wild-type P. aeruginosa with an estA-negative mutant and a strain overexpressing estA from a plasmid. Surprisingly, we observed that an EstA-deficient mutant strain failed to produce rhamnolipid whereas the overexpression of estA significantly enhanced the rhamnolipid yield in the culture supernatant. Furthermore, we could demonstrate that EstA is required for swimming, swarming, and twitching of P. aeruginosa. These effects were found to depend on the catalytic activity of EstA. An enzymatically inactive EstA variant obtained by replacing the catalytic serine with alanine failed to complement the estA mutant phenotypes.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Bacterial strains and growth conditions. The strains and plasmids used in this study are listed in Table 1. Escherichia coli was routinely grown at 37°C in Luria-Bertani (LB) medium or on LB agar plates (53). P. aeruginosa was grown at 30°C in LB medium or in rhamnolipid-promoting PPGAS medium (0.02 M NH₄Cl₂, 0.02 M KCl, 0.12 M Tris-HCl, 1.6 mM MgSO₄, 0.5% [wt/vol] glucose, 1% peptone; adjusted to pH 7.2) (62). Agrobacterium tumefaciens was grown at 28°C in AB minimal medium (12) supplemented with 0.2% glucose (ABG).

View this table: [in this window] [in a new window]

TABLE 1. Bacterial strains, plasmids, and primers

When required, antibiotics were added to the media at the following concentrations: tetracycline (Tc), 25 µg ml^–1 (E. coli) or 100 µg ml^–1 (P. aeruginosa); ampicillin, 100 µg ml^–1 (E. coli); carbenicillin, 500 µg ml^–1 (P. aeruginosa); streptomycin (Sm) and spectinomycin (Sp), 25 µg ml^–1 each (E. coli) or 100 µg ml^–1 each (P. aeruginosa); chloramphenicol, 50 µg ml^–1 (E. coli) or 300 µg ml^–1 (P. aeruginosa); gentamicin, 30 µg ml^–1 (A. tumefaciens).

General DNA manipulations. Plasmid DNA was purified as described by Birnboim and Doly (6), followed by anion-exchange chromatography on QIAGEN GmbH (Hilden, Germany) tips. Recombinant DNA techniques were performed essentially as described by Sambrook et al. (53). Restriction endonuclease reactions and bacteriophage T4 DNA ligase treatments were done as recommended by the manufacturers. DNA fragments were analyzed on 0.4 to 1% (wt/vol) agarose gels.

Construction of an estA-negative mutant. The suicide plasmid pMEestA, which harbors a Sm^r/Sp^r cassette inserted into the estA gene, was transformed into E. coli S17-1. This strain was used to transfer the plasmid into P. aeruginosa by diparental mating (55). For counterselection of E. coli S17-1 donor cells from P. aeruginosa after conjugation, Irgasan was used at a concentration of 25 µg ml^–1. The resulting Sm/Sp-resistant P. aeruginosa clones were tested for the loss of Tc resistance as an indication for successful allelic exchange of estA into estA::-Sm/Sp. Enrichment for Tc-sensitive cells was done using cycloserine at a concentration of 1.0 mg ml^–1 and Tc at 20 µg ml^–1 Successful inactivation was verified by PCR, Western blot analysis, and esterase activity assays.

Construction of an inactive EstA variant by site-directed mutagenesis. In order to inactivate the enzyme activity of the esterase, we changed the codon of active-site serine into an alanine coding triplet by overlap extension PCR (22) using the mutagenesis primers listed in Table 1. The resulting fragment was used to replace an internal EcoNI fragment of the wild-type estA gene in pBBX+, generating pBBXmut. The mutation was verified by DNA sequencing (QIAGEN GmbH, Hilden, Germany).

Overexpression and purification of EstA for antibody generation. Overexpression was done using pBR22estA in E. coli BL21(DE3). This plasmid is a derivative of pBR22b into which the open reading frame of the estA gene was inserted at the NdeI/XhoI sites. Expression was induced by addition of IPTG (isopropyl-ß-D-thiogalactopyranoside; 0.4 mM) at an optical density at 500 nm (OD₅₈₀) of 0.5, and cells were harvested 2 h after induction. EstA accumulated in inclusion bodies, which were collected by centrifugation of cell extracts after sonification of the cells. Chemically denatured (8 M urea) EstA was partly purified by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (5% stacking and 10% separating gels) (34) and used for the generation of a rabbit polyclonal anti-EstA serum (Eurogentec, Belgium). The specificity of the polyclonal serum was tested and found to be ideal at a dilution of 1:85,000.

Esterase assay. Cells harvested by centrifugation were washed in KP_i buffer (100 mM, pH 7.2) and disrupted by sonification, and the resulting cell extract was used to determine the esterase activity. The substrate (25 µl p-nitrophenyl-caproate [Sigma] dissolved in 5 ml ethanol) was added to 100 ml of KP_i (100 mM)-MgSO₄ (10 mM) buffer (pH 7.2). This test solution (1.5 ml) and cell extracts (5 to 20 µl) were mixed, and OD₄₁₀ min^–1 was monitored at 30°C.

Cultivation of P. aeruginosa biofilms. Biofilms were grown on glass slides (Superior; Marienfeld Company) in flow chambers with individual channel dimensions of 3 by 8 by 54 mm for 72 h at 30°C in modified PPGAS medium (0.05% glucose, 0.1% peptone, 2 mM NH₄Cl, 2 mM KCl, 100 mM Tris, 10 mM MgSO₄, 10 mM KNO₃, pH 7.5) or modified alginate-promoting medium (100 ml of a solution containing 100 mM Na-D-gluconate, 100 ml of a solution containing 100 mM KNO₃, and 50 ml of a solution containing 10 mM MgSO₄ and 750 ml phosphate buffer [1.03 g NaH₂PO₄, 2.93 g K₂HPO₄, pH 7.5]). All strains used for biofilm formation assays were chromosomally tagged with gfp as a fluorescent label for confocal laser scanning microscopy (31). The flow system was assembled and prepared as described previously (13). The cultures for inoculation of the flow chambers were prepared as follows. Overnight liquid cultures of P. aeruginosa were plated on PPGAS agar plates including the appropriate antibiotic and incubated overnight at 30°C. Thereafter bacteria from single colonies were resuspended in medium, cells were counted in a Abbe-Zeiss counting cell chamber, and the flow chambers were inoculated by injecting 1 x 10⁷ cells/ml in 5 ml flow medium. After inoculation flow channels were left without flow for 1 h for initial adhesion of the inoculum to the glass surface. Medium flow was kept at a constant rate of 20 ml/h, equivalent to a mean flow velocity of 0.02 mm/s.

Microscopy and image acquisition. All microscopic observations and image acquisitions were performed on a confocal laser scanning microscope (LSM 510; Carl Zeiss Jena, Jena, Germany) equipped with detectors and filter sets for monitoring green fluorescent protein fluorescence.

Motility. Surface motility of P. aeruginosa was monitored on PPGAS agar plates, which had agar concentrations of 0.5% for the swarm plates, 0.3% for the swim plates, and 1.5% for the twitching motility plates. The swarm agar plates contained glutamate (0.05%) instead of NH₄Cl. In the case of swimming and swarming the bacteria were staked on top of the agar, whereas for monitoring of twitching motility the bacteria where staked through the agar to colonize the interphase between the petri dish and agar.

Rhamnolipid analysis. Cultures were adjusted to an initial OD₅₈₀ of 0.05 and grown in 200 ml PPGAS medium supplemented with the appropriate antibiotics in 2-liter Erlenmeyer flasks at 30°C and 200 rpm for 48 h.

The orcinol assay is a colorimetric test to determine the amount of hexose sugars. This assay was used for the quantification of rhamnolipid (3-deoxy-hexose) in the culture supernatant. A volume of the culture supernatant (10 to 300 µl) was diluted with water to attain a volume of 300 µl, which was extracted twice with 600 µl diethylether. The pooled ether fractions were evaporated to dryness, and the remainder was dissolved in 100 µl distilled water and mixed with 100 µl 1.6% orcinol and 800 µl of 60% sulfuric acid. After heating to 80°C and shaking at 175 rpm for 30 min, the OD₄₂₁ was measured. The content of rhamnose in the samples was determined by comparing with rhamnose standards with a defined concentration. Rhamnolipid concentration was calculated based on the assumption that 1 µg of rhamnose corresponds to 2.5 µg of rhamnolipid (43).

TLC. The extracted rhamnolipids were dissolved in chloroform-methanol (9:1) in the concentration of 10 mg/ml, and 2 µl of the sample was spotted onto thin-layer chromatography (TLC) plates (silica gel 60 F₂₅₄; Merck). After development in chloroform-methanol-acetic acid (65:15:2), the TLC plates were dipped in the reagent 15% H₂SO₄ in ethanol and incubated for 2 to 5 min at 100°C.

Acyl-homoserine lactone (AHL) detection by an A. tumefaciens-based bioassay (9). Supernatants from P. aeruginosa stationary-phase cultures (500 µl) were extracted twice with 400 µl dichloromethane and dissolved in 5 µl of ethylacetate. Wells were punched in 1.5% agar plates with a sterile inoculating loop, and the extracts were filled in. After 15 min the ethylacetate was evaporated. Afterwards the plates were overlaid with 5 ml overnight culture of A. tumefaciens NTL4(pZLR4) in soft agar (soft ABG agar [0.7%] containing 40 µg 5-bromo-4-chloro-3-indolyl-ß-D-galactopyranoside [X-Gal]). The plates were incubated overnight at 30°C.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Phenotypic characterization of an estA-negative P. aeruginosa mutant. An estA-negative mutant of P. aeruginosa PAO1 was constructed by deletion of an internal fragment of the estA structural gene and insertion of a resistance marker cassette. This construct was then cloned into a suicide vector, and the resulting plasmid was used for homologous recombination, resulting in allelic exchange of estA in P. aeruginosa. As expected the estA mutant showed no esterase activity in preparations of the outer membrane, and Western blot analysis demonstrated that the EstA protein was completely absent. Preliminary characterization of the P. aeruginosa esterase-negative mutant revealed no obvious physiological differences compared to the wild type concerning the growth behavior under different culture conditions and in different growth media (data not shown). This indicates a physiological role for EstA which is not essential under standard laboratory conditions. For complementation purposes and moderate overexpression of the wild-type estA gene in P. aeruginosa the broad-host-range plasmid pBBX+ (63) was used.

EstA is required for rhamnolipid production. EstA has been shown to possess esterase activity, but the natural substrate of the enzyme is presently not known as well as the physiological circumstances under which its function is relevant for P. aeruginosa. However, it was not unrealistic that EstA, as a surface-exposed esterase, might be somehow involved in hydrolysis of ester bonds in compounds present on the cell surface or in the culture medium. Taking into account that EstA, as an autotransporter protein, was suspected of having virulence-related functions, we tentatively suspected rhamnolipids of being potential substrates for EstA because rhamnolipids are virulence determinants secreted by P. aeruginosa. The major rhamnolipids of P. aeruginosa contain one or two rhamnose moieties linked to two units of decanoic acids connected by an ester bond and thus might be cleavable by EstA.

In order to test the assumption that EstA might somehow be involved in the turnover or modification of rhamnolipid, its extracellular concentration was analyzed. Therefore, cultures of wild-type P. aeruginosa PAO1 and the estA mutant were grown in PPGAS medium, which is known to promote rhamnolipid production (62). Culture supernatants were collected after 48 h of growth, and rhamnolipids were extracted and subjected to TLC analysis. Indeed, we could detect a clear difference in rhamnolipid production between the two strains. As shown in Fig. 1 and 2 the amounts of both mono- and dirhamnolipid were affected in the estA mutant compared to the wild type. Unexpectedly, the estA mutant was completely unable to produce both major rhamnolipids (Fig. 2), which is contradictory to the assumption that inactivation of a potential rhamnolipid-hydrolyzing enzyme should result in accumulation of this compound. However, these findings suggested a potential role for EstA in rhamnolipid production rather than in its turnover by hydrolysis although its physiological role in this process was not obvious. The biosynthesis pathway of rhamnolipid is largely characterized, and there is no obvious requirement for an esterase activity in this pathway (56). At present it is completely unknown how rhamnolipids are secreted into the environment, and it can be speculated that this process may be mediated by a pore-forming protein of the outer membrane.

View larger version (7K): [in this window] [in a new window]

FIG. 1. Amounts of rhamnolipids in culture supernatants of different P. aeruginosa PAO1 strains. The amounts of rhamnolipids in culture supernatants were determined by an indirect assay (orcinol test). Wild-type (wt) P. aeruginosa PAO1, the estA mutant, and mutants overexpressing estA from plasmid pBBX+ and pBBXmut (inactive variant) were compared, and the rhlA mutant and strains containing the empty vector pBBR1MCS (V) served as controls. Culture supernatants were collected after 48 h of growth. The results were plotted into two diagrams because of the significant differences in the rhamnolipid amounts. Standard deviations were derived from three independent experiments.

View larger version (65K): [in this window] [in a new window]

FIG. 2. TLC separation of extracellular and cell-bound rhamnolipids from P. aeruginosa. P. aeruginosa PAO1 (wild type [wt]) and the estA-negative mutant (estA-) were examined for rhamnolipid production. Rhamnolipids were extracted from cultures which were grown for 48 h in PPGAS medium. The rhamnolipid standard (RL) was purified from P. aeruginosa. Rhamnolipids were separated by TLC and stained with a reagent containing orcinol and sulfuric acid in ethanol.

EstA belongs to the autotransporter protein family and is composed of two distinct domains, of which the N-terminal domain harbors the enzymatic activity whereas the C-terminal transporter domain represents a beta-barrel pore in the outer membrane. Although it has not been reported that substrates other than the corresponding passenger domain itself can be secreted across the outer membrane by these domains, one can speculate that the pore-forming domain of the EstA transporter domain might serve as a translocation pore for rhamnolipid.

In order to distinguish whether the pore-forming C-terminal translocator domain of EstA might be the relevant part of EstA for rhamnolipid production and responsible for the rhamnolipid-negative phenotype of the estA mutant rather than the enzyme activity of the passenger domain, we constructed an inactive variant of EstA. The predicted active-site serine₃₈ residue (63) was replaced by an alanine using site-directed mutagenesis. The loss of enzyme activity of the EstA variant (EstA*) was confirmed by expression from plasmid pBBXmut in the estA-negative mutant and subsequent activity measurement, thereby demonstrating that in fact serine₃₈ is the active-site serine. In addition, expression of EstA* and correct localization of the protein in the membrane were verified by Western blot analysis and found to be unaffected by the mutation (data not shown). In addition we know also from our work that even in the heterologous host E. coli another inactive EstA* variant resides on the cell surface (5).

The amount of rhamnolipids produced by the strains was determined by the orcinol assay (10). The cultures were grown as described above; culture supernatants were collected, and rhamnolipids were extracted. The data were corrected according to the minimal variation of cell densities. The results of these experiments shown in Fig. 1 revealed a significant difference in the amounts of rhamnolipids between wild-type, estA-negative, and estA-overexpressing strains. The rhlA-negative strain is known to be completely defective in rhamnolipid synthesis and thus served as a negative control. The rhamnolipid production of the estA mutant could be restored by complementation with the plasmid pBBX+, harboring the wild-type estA gene. In contrast, complementation failed when the inactive estA variant was expressed from the same plasmid (pBBXmut). Interestingly pBBX+ not only restored rhamnolipid production of the mutant strain but also significantly increased the amount of rhamnolipid produced when it was introduced into the wild type. These data indicate that rhamnolipid production of P. aeruginosa essentially depends on EstA, which appears to play the role in rhamnolipid biosynthesis of limiting the yield of rhamnolipids in the wild-type background. Moreover, the failure of the inactive variant in complementation of EstA-dependent rhamnolipid production demonstrates that the enzymatic activity of EstA in fact is the relevant function of the protein for rhamnolipid production. This is consistent with the finding that no accumulation of rhamnolipids within the cell was observed in the estA mutant (Fig. 2).

The production of rhamnolipid in P. aeruginosa is regulated by a quorum-sensing system involving AHLs as signal molecules or "autoinducers." Riedel and coworkers (51) found that inactivation of an EstA homologue in Serratia liquefaciens resulted in reduced levels of AHLs. However, this effect was observed only under specific artificial experimental conditions, namely, when the growth medium was supplemented with unusually high concentrations of Tween 20 as artificial lipid substrates, whereas it remained uncertain without lipid supplementation (51). In the experiments presented in this report the bacteria were grown without any lipid supplementation. Under these conditions the estA-negative mutant produced signal molecules which could be detected by the A. tumefaciens-based bioassay (Fig. 3). In this bioassay AHLs are detected using a reporter strain which harbors an AHL-responsive promoter fused to the ß-galactosidase gene (lacZ::traG), which converts X-Gal, resulting in formation of blue halos in the presence of autoinducers (9). Compared to supernatants of the wild type, those of the estA-negative mutant raised halos of the same size and thus contained equal amounts of AHLs. In contrast supernatants of the lasRI rhlRI quorum-sensing mutant, which served as a negative control, produced no visible halo in the bioassay. This demonstrates that the production of the AHLs detected by the A. tumefaciens bioassay is unaffected in the estA mutant and that the EstA-dependent effects on rhamnolipid production are not the result of alterations of these AHL levels.

View larger version (101K): [in this window] [in a new window]

FIG. 3. Agrobacterium tumefaciens-based bioassay for autoinducer (AI) detection in culture supernatants of P. aeruginosa. Tested were wild-type (WT) P. aeruginosa PAO1 and the estA-negative mutant for the presence of AIs, and an AI-deficient mutant (rhlRI and lasRI deficient) served as a negative control. Shown is the A. tumefaciens-based bioassay, where the formation of blue halos confirmed the presence of AI molecules.

Motility and biofilm formation of the estA mutant. Some phenotypes of P. aeruginosa which are physiologically connected to rhamnolipid production were investigated in the estA mutant. Rhamnolipid production has been reported to promote surface translocation on semisolid surfaces, i.e., swarming motility is facilitated by reduced friction through the action of rhamnolipid detergent (32). In fact, as expected, the esterase-negative mutant strain showed no swarming motility (Fig. 4), which is consistent with the lack of rhamnolipid production.

View larger version (68K): [in this window] [in a new window]

FIG. 4. Swimming (A), swarming (B), and twitching (C) motility of P. aeruginosa PAO1. Wild-type P. aeruginosa (wt) and estA-negative mutant (estA-) strains were analyzed on agar plates. In addition the estA-negative mutant was transformed with different plasmids, i.e., the empty vector (v) as a negative control, the estA-carrying plasmid pBBX+, and a mutated version of pBBX coding for an inactive esterase (pBBXmut). The agar plates contained different agar concentrations to establish swimming (0.3%) and swarming (0.5%) motility and were incubated overnight at room temperature (swimming) or at 30°C (swarming plus twitching). (D) Motility of the P. aeruginosa estA mutant with the addition of exogenous rhamnolipids (RLs) and the wild type was monitored. Extracted RL was added to the agar plates at a concentration of 0.4 g/liter, which represents the wild-type level of RLs.

Swarming motility of P. aeruginosa depends on the functionality of both type IV pili and flagella (32). To further dissect the swarming motility deficiency phenotype of the EstA mutant, twitching motility and swimming were also analyzed. Interestingly, in addition to swarming, both other forms of surface motility were completely absent in the estA mutant. Motility could not be restored by addition of wild-type amounts of rhamnolipid to the growth medium of the estA mutant, suggesting that the rhamnolipid deficiency is not the direct cause of the nonmotile phenotype (Fig. 4D).

In biofilm formation the ability of bacterial cells to move on surfaces plays an important role in achieving the mature three-dimensional biofilm architecture (29, 30). Since the esterase mutant strain appeared to be unable to display any type of motility, we suspected that the formation or the structure of biofilms might be different from that of wild-type biofilms.

Indeed, differences in biofilm formation and maintenance were observed with this strain. Compared to that of the wild type, the estA mutant biofilms were characterized by a significant reduction in surface coverage. The mutant strain appeared to form biofilms consisting of large aggregates of cells with regions of extremely low cell densities between the individual microcolonies, as expected for nonmotile strains that can expand only by clonal growth (Fig. 5).

View larger version (49K): [in this window] [in a new window]

FIG. 5. Biofilm confocal laser scanning micrographs of different P. aeruginosa strains. P. aeruginosa PAO1, the estA mutant, and mutants overexpressing estA from plasmids pBBX+ and pBBXmut (inactive variant) were compared, and the strain containing the empty vector pBBR1MCS (V) served as a control. Biofilms were grown in a flow cell, and section pictures were taken after 24 and 72 h of growth with x630 magnification.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
We have previously identified a lipolytic enzyme located in the outer membrane of P. aeruginosa. This enzyme was designated esterase EstA and was shown to be exposed on the surface (63). The esterase is secreted to the outer surface by the so-called autotransporter mechanism or type V secretion pathway (21, 63). After translocation to the cell surface, most autotransporter proteins are processed proteolytically and the N-terminal passenger domain is released from the outer membrane or remains attached to the translocator domain by noncovalent interactions. There are very few exceptions, i.e., where autotransporter passenger domains remain covalently attached to their translocator domains. However, most of these autotransporter proteins belong to the "oca" subfamily of trimeric adhesins (23, 65). In contrast, McaP from Moraxella catarrhalis, EstA from Serratia liquefaciens, and P. aeruginosa EstA represent prototype proteins of a unique family of autotransporter proteins in which the passenger domain carrying the enzyme activity remains covalently attached to the translocator domain (51, 59, 63). Whereas an adhesin remaining bound to the cell surface can easily be considered a biological benefit, the physiological need for a covalently cell surface-bound lipolytic activity, as for EstA or McaP, is less obvious. However, this unique surface exposition of these enzymes in combination with the generally virulence-associated role of autotransporter proteins suggested that the physiological role of these cell surface esterases might be somehow related to the turnover of lipid compounds, which may be extracellular or derived from the outer membrane. This led us to the assumption that rhamnolipids as extracellular lipid compounds which contain ester bonds might be potential substrates for EstA. Since rhamnolipids are also virulence factors, this would create a link between EstA and virulence. In order to investigate the potential effect of EstA on the yield of rhamnolipids in P. aeruginosa, an EstA-deficient mutant was constructed, and this strain was found to produce only marginal amounts of rhamnolipid. This was unexpected, since it suggests that EstA plays a potential role in rhamnolipid production, although there is no obvious function in the rhamnolipid biosynthesis pathway which might directly require the enzymatic activity of an esterase. Rhamnolipid production in P. aeruginosa is controlled by the hierarchical quorum-sensing regulatory cascade composed oft the LasR/I and RhlR/I systems (56). In S. liquefaciens EstA has been suspected of indirectly influencing the synthesis of quorum-sensing molecules by providing the cells with fatty acids (51). However, this was only the case when cells were grown in media supplemented with high concentrations of Tween 20 as a synthetic lipid substrate. Without lipid supplementation S. liquefaciens EstA had no effect on autoinducer synthesis and the relevance of the observed effects on autoinducer production under physiological conditions remained unclear (51). In contrast to this the growth medium used in our study was not supplemented with lipids as potential substrates for EstA of P. aeruginosa. The A. tumefaciens bioassay for autoinducers of the AHL type revealed that the production of the detectable signal molecules was not affected in the estA mutant under these growth conditions. This indicates that the observed decrease in rhamnolipid production of the EstA-negative strain is probably not due to the level of autoinducer molecules and the corresponding effects on quorum-sensing regulation.

Inactivation of the estA gene not only resulted in rhamnolipid deficiency but also influenced other virulence-related functions like cellular motility, i.e., swimming, twitching motility, and swarming. So far, among the three modes of motility of P. aeruginosa only swarming has been found to depend on the presence of rhamnolipid, probably due to the wetting properties of this biodetergent, which might be required for the movement of cells on semisolid surfaces (16, 32). A physiological link between swimming and twitching motility has not been described so far, although both are well characterized. This strongly suggests that the decreased amount of rhamnolipids in the estA mutant is not the reason for these motility defects but is an additional phenotype which is somehow coinfluenced by the inactivation of EstA.

Consistent with the finding that rhamnolipid production and motility were severely affected by EstA, biofilms formed by the estA mutant were also different from wild-type biofilms. The formation of biofilms in P. aeruginosa is a highly complex process in which motility of the cells, nutrition, quorum sensing, and rhamnolipid production play important roles to obtain the architecture of mature biofilms (15, 29, 30).

The capability of cellular motility and rhamnolipid production could be restored when the wild-type gene was expressed from a plasmid, whereas the inactive variant was not functional for complementation of these phenotypes. The loss of rhamnolipid production in the estA mutant could serve as only a partial explanation for the observed motility phenotypes in the case of swarming. Interestingly, this is consistent with the finding that motility of the EstA mutant could not be restored by the exogenous addition of rhamnolipids to the medium (Fig. 2). Moreover, the wild type and the estA mutant were mixed in equal quantities and cocultivated on swarming assay plates, and cells were isolated from the swarming front. As assayed by determining the CFU on antibiotic agar plates only 0.1% of the cells in the swarming front were of the estA mutant strain (data not shown). This strongly suggests that swarming cannot be rescued for the estA mutant by any diffusible compound produced by EstA in the wild type. The fact that the enzyme activity of EstA is responsible for the observed phenotypes shows that, probably by hydrolysis of a yet-unknown compound, EstA stimulates rhamnolipid production, cell motility, and possibly other yet-unrevealed cellular functions. Preliminary results of transcriptome analysis after 6 and 24 h of growth suggest a set of possible molecular explanations for single effects observed with the estA mutant (data not shown). Consistent with the rhamnolipid deficiency, the genes rhlB and rhlC, coding for rhamnosyltransferases I and II, the key enzymes in rhamnolipid biosynthesis, appear to be significantly downregulated in the estA mutant. In addition, expression of several genes involved in the biogenesis of type IV pili is reduced in the mutant. Among them are the pilA, pilX, pilG, and pilB genes, which are encoded in different operons at different loci of the P. aeruginosa chromosome. This strongly suggests that pilus biosynthesis and function are severely affected in the estA mutant, which in turn may explain the defects in twitching and swarming motility and, in part, the differences in biofilm architecture. In contrast, there is no indication of different expression of genes involved in biosynthesis or function of flagella which could serve as an explanation for the lack of swimming. The different phenotypes affected by EstA have no obvious physiological or regulatory link and thus cannot be explained satisfyingly and in a comprehensive way at the moment. Myxococcus xanthus and P. aeruginosa exhibit type IV pilus-dependent cellular motility, and both are affected by gradients of phospholipids as a kind of chemoattractant (8, 27, 28). In P. aeruginosa this chemotaxis-like motility has been shown to depend on the activity of an extracellular phospholipase C, which liberates diacylglycerol from phosphatidylethanolamine (2). This reaction is supposed to be the first step in a potential lipid signaling-like process which may involve further reactions catalyzed by other lipolytic enzymes like esterases and lipases (8). EstA has to be considered a candidate enzyme for such downstream reactions. However, not only twitching motility is affected by EstA; all other forms of motility and cellular functions like rhamnolipid production and biofilm formation appear to require this enzyme. This indicates that a potential signal molecule generated by the EstA enzyme activity may be part of an important signaling cascade which may even be involved in the regulation or modulation of other virulence-associated functions of P. aeruginosa. However, to define an exact role of EstA will require analysis of the lipid composition as well as further detailed analysis of transcriptome data.

 
We thank S. Heeb, V. Wright, M. Camara, and P. Williams (Institute of Infection Immunity and Inflammation and School of Molecular Medical Sciences, University of Nottingham, United Kingdom) for help with preliminary transcriptome analysis and excellent discussions.

Work of S. Wilhelm is supported by the DFG framework of the Schwerpunktprojekt 1170: Directed Evolution To Optimize and Understand Molecular Biocatalysis.

 
* Corresponding author. Mailing address: Institute for Molecular Enzyme Technology, Heinrich Heine University Duesseldorf, Research Centre Juelich, Stetternicher Forst, D-52426 Juelich, Germany. Phone: 492461612947. Fax: 492461612490. E-mail: f.rosenau{at}fz-juelich.de

Published ahead of print on 13 July 2007.

Present address: Institute for Microbiology, Technical University Braunschweig, D-38023 Braunschweig, Germany.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 

1
1. Al-Tahhan, R. A., T. R. Sandrin, A. A. Bodour, and R. M. Maier. 2000. Rhamnolipid-induced removal of lipopolysaccharide from Pseudomonas aeruginosa: effect on cell surface properties and interaction with hydrophobic substrates. Appl. Environ. Microbiol. 66:3262-3268.[Abstract/Free Full Text]
2
2. Barker, A. P., A. I. Vasil, A. Filloux, G. Ball, P. J. Wilderman, and M. L. Vasil. 2004. A novel extracellular phospholipase C of Pseudomonas aeruginosa is required for phospholipid chemotaxis. Mol. Microbiol. 53:1089-1098.[CrossRef][Medline]
3
3. Beal, R., and W. B. Betts. 2000. Role of rhamnolipid biosurfactants in the uptake and mineralization of hexadecane in Pseudomonas aeruginosa. J. Appl. Microbiol. 89:158-168.[CrossRef][Medline]
4
4. Beatson, S. A., C. B. Whitchurch, A. B. Semmler, and J. S. Mattick. 2002. Quorum sensing is not required for twitching motility in Pseudomonas aeruginosa. J. Bacteriol. 184:3598-3604.[Abstract/Free Full Text]
5
5. Becker, S., S. Theile, N. Heppeler, A. Michalczyk, A. Wentzel, S. Wilhelm, K. E. Jaeger, and H. Kolmar. 2005. A generic system for the Escherichia coli cell-surface display of lipolytic enzymes. FEBS Lett. 579:1177-1182.[CrossRef][Medline]
6
6. Birnboim, H. C., and J. Doly. 1979. A rapid alkaline extraction procedure for screening recombinant plasmid DNA. Nucleic Acids Res. 7:1513-1523.[Abstract/Free Full Text]
7
7. Boles, B. R., M. Thoendel, and P. K. Singh. 2005. Rhamnolipids mediate detachment of Pseudomonas aeruginosa from biofilms. Mol. Microbiol. 57:1210-1223.[CrossRef][Medline]
8
8. Bonner, P. J., and L. J. Shimkets. 2006. Phospholipid directed motility of surface-motile bacteria. Mol. Microbiol. 61:1101-1109.[CrossRef][Medline]
9
9. Cha, C., P. Gao, Y.-C. Chen, P. D. Shaw, and S. K. Farrand. 1998. Production of acyl-homoserine lactone quorum-sensing signals by Gram-negative plant-associated bacteria. Mol. Plant-Microbe Interact. 11:1119-1129.[Medline]
10
10. Chandrasekaran, E. V., and J. N. Bemiller. 1980. Constituent analyses of glycosaminoglycans, p. 89-96. In R. L. Whistler and J. N. Bemiller (ed.), Methods in carbohydrate chemistry. Academic Press, New York, NY.
11
11. Chen, G., and H. Zhu. 2005. lux-marked Pseudomonas aeruginosa lipopolysaccharide production in the presence of rhamnolipid. Colloids Surf. 41:43-48.[CrossRef]
12
12. Chilton, M. D., T. C. Currier, S. K. Farrand, A. J. Bendich, M. P. Gordon, and E. W. Nester. 1974. Agrobacterium tumefaciens DNA and PS8 bacteriophage DNA not detected in crown gall tumors. Proc. Natl. Acad. Sci. USA 71:3672-3676.[Abstract/Free Full Text]
13
13. Christensen, B. B., C. Sternberg, J. B. Andersen, R. J. Palmer, Jr., A. T. Nielsen, M. Givskov, and S. Molin. 1999. Molecular tools for study of biofilm physiology. Methods Enzymol. 310:20-42.[Medline]
14
14. Cosson, P., L. Zulianello, O. Join-Lambert, F. Faurisson, L. Gebbie, M. Benghezal, C. Van Delden, L. K. Curty, and T. Kohler. 2002. Pseudomonas aeruginosa virulence analyzed in a Dictyostelium discoideum host system. J. Bacteriol. 184:3027-3033.[Abstract/Free Full Text]
15
15. Davey, M. E., N. C. Caiazza, and G. A. O'Toole. 2003. Rhamnolipid surfactant production affects biofilm architecture in Pseudomonas aeruginosa PAO1. J. Bacteriol. 185:1027-1036.[Abstract/Free Full Text]
16
16. Deziel, E., F. Lepine, S. Milot, and R. Villemur. 2003. rhlA is required for the production of a novel biosurfactant promoting swarming motility in Pseudomonas aeruginosa: 3-(3-hydroxyalkanoyloxy)alkanoic acids (HAAs), the precursors of rhamnolipids. Microbiology 149:2005-2013.[Abstract/Free Full Text]
17
17. Reference deleted.
18
18. Espinosa-Urgel, M. 2003. Resident parking only: rhamnolipids maintain fluid channels in biofilms. J. Bacteriol. 185:699-700.[Free Full Text]
19
19. Haba, E., A. Pinazo, O. Jauregui, M. J. Espuny, M. R. Infante, and A. Manresa. 2003. Physicochemical characterization and antimicrobial properties of rhamnolipids produced by Pseudomonas aeruginosa 47T2 NCBIM 40044. Biotechnol. Bioeng. 81:316-322.[CrossRef][Medline]
20
20. Henderson, I. R., and J. P. Nataro. 2001. Virulence functions of autotransporter proteins. Infect. Immun. 69:1231-1243.[Free Full Text]
21
21. Henderson, I. R., F. Navarro-Garcia, M. Desvaux, R. C. Fernandez, and D. Ala'Aldeen. 2004. Type V protein secretion pathway: the autotransporter story. Microbiol. Mol. Biol. Rev. 68:692-744.[Abstract/Free Full Text]
22
22. Ho, S. N., H. D. Hunt, R. M. Horton, J. K. Pullen, and L. R. Pease. 1989. Site-directed mutagenesis by overlap extension using the polymerase chain reaction. Gene 77:51-59.[CrossRef][Medline]
23
23. Hoiczyk, E., A. Roggenkamp, M. Reichenbecher, A. Lupas, and J. Heesemann. 2000. Structure and sequence analysis of Yersinia YadA and Moraxella UspAs reveal a novel class of adhesins. EMBO J. 19:5989-5999.[CrossRef][Medline]
24
24. Holloway, B. W., V. Krishnapillai, and A. F. Morgan. 1979. Chromosomal genetics of Pseudomonas. Microbiol. Rev. 43:73-102.[Free Full Text]
25
25. Irie, Y., G. A. O'Toole, and M. H. Yuk. 2005. Pseudomonas aeruginosa rhamnolipids disperse Bordetella bronchiseptica biofilms. FEMS Microbiol. Lett. 250:237-243.[CrossRef][Medline]
26
26. Johnson, M., and D. Boese-Marrazzo. 1980. Production and properties of heat-stable extracellular hemolysin from Pseudomonas aeruginosa. Infect. Immun. 29:1028-1033.[Abstract/Free Full Text]
27
27. Kearns, D. B., J. Robinson, and L. J. Shimkets. 2001. Pseudomonas aeruginosa exhibits directed twitching motility up phosphatidylethanolamine gradients. J. Bacteriol. 183:763-767.[Abstract/Free Full Text]
28
28. Kearns, D. B., and L. J. Shimkets. 1998. Chemotaxis in a gliding bacterium. Proc. Natl. Acad. Sci. USA 95:11957-11962.[Abstract/Free Full Text]
29
29. Klausen, M., A. Aaes-Jorgensen, S. Molin, and T. Tolker-Nielsen. 2003. Involvement of bacterial migration in the development of complex multicellular structures in Pseudomonas aeruginosa biofilms. Mol. Microbiol. 50:61-68.[CrossRef][Medline]
30
30. Klausen, M., A. Heydorn, P. Ragas, L. Lambertsen, A. Aaes-Jorgensen, S. Molin, and T. Tolker-Nielsen. 2003. Biofilm formation by Pseudomonas aeruginosa wild type, flagella and type IV pili mutants. Mol. Microbiol. 48:1511-1524.[CrossRef][Medline]
31
31. Koch, B., L. E. Jensen, and O. Nybroe. 2001. A panel of Tn7-based vectors for insertion of the gfp marker gene or for delivery of cloned DNA into Gram-negative bacteria at a neutral chromosomal site. J. Microbiol. Methods 45:187-195.[CrossRef][Medline]
32
32. Kohler, T., L. K. Curty, F. Barja, C. van Delden, and J. C. Pechere. 2000. Swarming of Pseudomonas aeruginosa is dependent on cell-to-cell signaling and requires flagella and pili. J. Bacteriol. 182:5990-5996.[Abstract/Free Full Text]
33
33. Kovach, M. E., R. W. Phillips, P. H. Elzer, R. M. Roop II, and K. M. Peterson. 1994. pBBR1MCS: a broad-host-range cloning vector. BioTechniques. 16:800-802.[Medline]
34
34. Laemmli, U. K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680-685.[CrossRef][Medline]
35
35. Lepine, F., E. Deziel, S. Milot, and R. Villemur. 2002. Liquid chromatographic/mass spectrometric detection of the 3-(3-hydroxyalkanoyloxy) alkanoic acid precursors of rhamnolipids in Pseudomonas aeruginosa cultures. J. Mass Spectrom. 37:41-46.[CrossRef][Medline]
36
36. Luo, Z. Q., S. Su, and S. K. Farrand. 2003. In situ activation of the quorum-sensing transcription factor TraR by cognate and noncognate acyl-homoserine lactone ligands: kinetics and consequences. J. Bacteriol. 185:5665-5672.[Abstract/Free Full Text]
37
37. Maier, R. M., and G. Soberon-Chavez. 2000. Pseudomonas aeruginosa rhamnolipids: biosynthesis and potential applications. Appl. Microbiol. Biotechnol. 54:625-633.[CrossRef][Medline]
38
38. Matsuyama, T., and Y. Nakagawa. 1996. Bacterial wetting agents working in colonization of bacteria on surface environments. Colloids Surf. 7:207-214.[CrossRef]
39
39. Mattick, J. S. 2002. Type IV pili and twitching motility. Annu. Rev. Microbiol. 56:289-314.[CrossRef][Medline]
40
40. Reference deleted.
41
41. McClure, C. D., and N. L. Schiller. 1996. Inhibition of macrophage phagocytosis by Pseudomonas aeruginosa rhamnolipids in vitro and in vivo. Curr. Microbiol. 33:109-117.[CrossRef][Medline]
42
42. Noordman, W. H., and D. B. Janssen. 2002. Rhamnolipid stimulates uptake of hydrophobic compounds by Pseudomonas aeruginosa. Appl. Environ Microbiol. 68:4502-4508.[Abstract/Free Full Text]
43
43. Ochsner, U. A., A. K. Koch, A. Fiechter, and J. Reiser. 1994. Isolation and characterization of a regulatory gene affecting rhamnolipid biosurfactant synthesis in Pseudomonas aeruginosa. J. Bacteriol. 176:2044-2054.[Abstract/Free Full Text]
44
44. Parales, R. E., and C. S. Harwood. 1993. Construction and use of a new broad-host-range lacZ transcriptional fusion vector, pHRP309, for gram^– bacteria. Gene 29:23-30.
45
45. Potvin, E., D. E. Lehoux, I. Kukavica-Ibrulj, K. L. Richard, F. Sanschagrin, G. W. Lau, and R. C. Levesque. 2003. In vivo functional genomics of Pseudomonas aeruginosa for high-throughput screening of new virulence factors and antibacterial targets. Environ. Microbiol. 5:1294-1308.[CrossRef][Medline]
46
46. Rahim, R., U. A. Ochsner, C. Olvera, M. Graninger, P. Messner, J. S. Lam, and G. Soberon-Chavez. 2001. Cloning and functional characterization of the Pseudomonas aeruginosa rhlC gene that encodes rhamnosyltransferase 2, an enzyme responsible for di-rhamnolipid biosynthesis. Mol. Microbiol. 40:708-718.[CrossRef][Medline]
47
47. Rahme, L. G., F. M. Ausubel, H. Cao, E. Drenkard, B. C. Goumnerov, G. W. Lau, S. Mahajan-Miklos, J. Plotnikova, M. W. Tan, J. Tsongalis, C. L. Walendziewicz, and R. G. Tompkins. 2000. Plants and animals share functionally common bacterial virulence factors. Proc. Natl. Acad. Sci. USA 97:8815-8821.[Abstract/Free Full Text]
48
48. Rashid, M. H., and A. Kornberg. 2000. Inorganic polyphosphate is needed for swimming, swarming, and twitching motilities of Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. USA 97:4885-4890.[Abstract/Free Full Text]
49
49. Read, R. C., P. Roberts, N. Munro, A. Rutman, A. Hastie, T. Shryock, R. Hall, W. McDonald-Gibson, V. Lund, G. Taylor, et al. 1992. Effect of Pseudomonas aeruginosa rhamnolipids on mucociliary transport and ciliary beating. J. Appl. Physiol. 72:2271-2277.[Abstract/Free Full Text]
50
50. Rendell, N. B., G. W. Taylor, M. Somerville, H. Todd, R. Wilson, and P. J. Cole. 1990. Characterisation of Pseudomonas rhamnolipids. Biochim. Biophys. Acta 1045:189-193.[Medline]
51
51. Riedel, K., D. Talker-Huiber, M. Givskov, H. Schwab, and L. Eberl. 2003. Identification and characterization of a GDSL esterase gene located proximal to the swr quorum-sensing system of Serratia liquefaciens MG1. Appl. Environ Microbiol. 69:3901-3910.[Abstract/Free Full Text]
52
52. Rosenau, F., and K.-E. Jaeger. 2004. Overexpression and secretion of Pseudomonas lipases, p. 491-508. In J. L. Ramos (ed.), Pseudomonas, vol. 3. Kluwer Academic/Plenum Publishers, New York, NY.
53
53. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
54
54. Sharma, M., and S. Anand. 2002. Swarming: a coordinated bacterial activity. Curr. Sci. 83:707-718.
55
55. Simon, R., U. Priefer, and A. Pühler. 1983. A broad host range mobilization system for in vitro genetic engineering: transposon mutagenesis in Gram-negative bacteria. Bio/Technology 1:784-791.[CrossRef]
56
56. Soberon-Chavez, G., F. Lepine, and E. Deziel. 2005. Production of rhamnolipids by Pseudomonas aeruginosa. Appl. Microbiol. Biotechnol. 14:1-8.
57
57. Studier, F. W., and B. A. Moffatt. 1986. Use of bacteriophage T7 RNA polymerase to direct selective high level expression of cloned genes. J. Mol. Biol. 189:113-130.[CrossRef][Medline]
58
58. Studier, F. W. 1991. Use of bacteriophage T7 lysozyme to improve an inducible T7 expression system. J. Mol. Biol. 219:37-44.[CrossRef][Medline]
59
59. Timpe, J. M., M. M. Holm, S. L. Vanlerberg, V. Basrur, and E. R. Lafontaine. 2003. Identification of a Moraxella catarrhalis outer membrane protein exhibiting both adhesin and lipolytic activities. Infect. Immun. 71:4341-4350.[Abstract/Free Full Text]
60
60. Voisard, C., C. T. Bull, C. Keel, J. Laville, M. Maurhofer, U. Schnider, G. De'fago, and D. Haas. 1994. Biocontrol of root diseases by Pseudomonas fluorescens CHA0: current concepts and experimental approaches, p. 67-89. In F. O'Gara, D. N. Dowling, and B. Boesten (ed.), Molecular ecology of rhizosphere microorganisms. VCH, Weinheim, Germany.
61
61. Watson, A. A., R. A. Alm, and J. S. Mattick. 1996. Construction of improved vectors for protein production in Pseudomonas aeruginosa. Gene 172:163-164.[CrossRef][Medline]
62
62. Wild, M., A. D. Caro, A. L. Hernandez, R. M. Miller, and G. Soberon-Chavez. 1997. Selection and partial characterization of a Pseudomonas aeruginosa mono-rhamnolipid deficient mutant. FEMS Microbiol. Lett. 153:279-285.[CrossRef][Medline]
63
63. Wilhelm, S., J. Tommassen, and K. E. Jaeger. 1999. A novel lipolytic enzyme located in the outer membrane of Pseudomonas aeruginosa. J. Bacteriol. 181:6977-6986.[Abstract/Free Full Text]
64
64. Yanisch-Perron, C., J. Vieira, and J. Messing. 1985. Improved M13 phage cloning vectors and host strains: nucleotide sequences of the M13mp18 and pUC19 vectors. Gene 33:103-119.[CrossRef][Medline]
65
65. Yeo, H. J., S. E. Cotter, S. Laarmann, T. Juehne, J. W. St Geme, and G. Waksman. 2004. Structural basis for host recognition by the Haemophilus influenzae Hia autotransporter. EMBO J. 23:1245-1256.[CrossRef][Medline]
66
66. Zhang, Y., and R. M. Miller. 1994. Effect of a Pseudomonas rhamnolipid biosurfactant on cell hydrophobicity and biodegradation of octadecane. Appl. Environ. Microbiol. 60:2101-2106.[Abstract/Free Full Text]

---

Journal of Bacteriology, September 2007, p. 6695-6703, Vol. 189, No. 18
0021-9193/07/$08.00+0     doi:10.1128/JB.00023-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

This article has been cited by other articles:

• Miller, R. M., Tomaras, A. P., Barker, A. P., Voelker, D. R., Chan, E. D., Vasil, A. I., Vasil, M. L. (2008). Pseudomonas aeruginosa Twitching Motility-Mediated Chemotaxis towards Phospholipids and Fatty Acids: Specificity and Metabolic Requirements. J. Bacteriol. 190: 4038-4049 [Abstract] [Full Text]  

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Wilhelm, S. Articles by Jaeger, K.-E. Search for Related Content PubMed PubMed Citation Articles by Wilhelm, S. Articles by Jaeger, K.-E.