• We are sorry, but NCBI web applications do not support your browser and may not function properly. More information
Logo of jbacterPermissionsJournals.ASM.orgJournalJB ArticleJournal InfoAuthorsReviewers
J Bacteriol. Mar 2007; 189(6): 2531–2539.
Published online Jan 12, 2007. doi:  10.1128/JB.01515-06
PMCID: PMC1899385

Multiple Roles of Biosurfactants in Structural Biofilm Development by Pseudomonas aeruginosa[down-pointing small open triangle]


Recent studies have indicated that biosurfactants produced by Pseudomonas aeruginosa play a role both in maintaining channels between multicellular structures in biofilms and in dispersal of cells from biofilms. Through the use of flow cell technology and enhanced confocal laser scanning microscopy, we have obtained results which suggest that the biosurfactants produced by P. aeruginosa play additional roles in structural biofilm development. We present genetic evidence that during biofilm development by P. aeruginosa, biosurfactants promote microcolony formation in the initial phase and facilitate migration-dependent structural development in the later phase. P. aeruginosa rhlA mutants, deficient in synthesis of biosurfactants, were not capable of forming microcolonies in the initial phase of biofilm formation. Experiments involving two-color-coded mixed-strain biofilms showed that P. aeruginosa rhlA mutants were defective in migration-dependent development of mushroom-shaped multicellular structures in the later phase of biofilm formation. Experiments involving three-color-coded mixed-strain P. aeruginosa biofilms demonstrated that the wild-type and rhlA and pilA mutant strains formed distinct subpopulations on top of each other dependent on their ability to migrate and produce biosurfactants.

Evidence is accumulating that many infections display elevated tolerance towards antimicrobial attack because the causative bacteria reside in biofilms (8, 15, 51). The opportunistic human pathogen Pseudomonas aeruginosa has become a model organism for studying biofilm development. When P. aeruginosa is grown as a biofilm in flow chambers, it often develops mushroom-shaped multicellular structures separated by liquid-filled channels (9, 26, 32). The cap-forming subpopulation and the stalk-forming subpopulation of these mushroom-shaped structures in many cases display differential tolerance to antimicrobial compounds. For example, the antibiotic tobramycin was shown to kill preferentially bacteria in the cap portion of the mushroom-shaped structures, whereas the antibiotic colistin, the detergent sodium dodecyl sulfate, and the chelator EDTA were shown to kill preferentially bacteria in the stalk portion of the mushroom-shaped structures (3, 5, 19, 21). Knowledge about subpopulation development in biofilms, and the way the subpopulations interact and change during structural biofilm development, may be useful for creating strategies to control biofilm formation and eradicate persistent infections.

Studies of liquid flow and molecular diffusion in flow chamber-grown biofilms have led to the proposal that the channels and interstitial voids between the microcolonies may function as a circulation system for efficient nutrient supply and waste product removal (11, 52). Biosurfactants produced by P. aeruginosa via the RhlA gene product were shown to be important for maintenance of the water channels between the mushroom-shaped multicellular structures in biofilms (10). A study which employed fluorescent reporter genes indicated that biosurfactant synthesis preferentially takes place in microcolonies during early stages of biofilm development and in the stalk portion of the mushroom-shaped structures in more mature P. aeruginosa biofilms (33). In addition, evidence has been presented that the biosurfactants produced by P. aeruginosa are involved in the dispersion of cells from biofilms (5, 24, 47).

P. aeruginosa produces a number of biosurfactants, of which the three most abundant are 3-(3-hydroxyalkanoyloxy)alcanoic acid (HAA), l-rhamnosyl-3-hydroxydecanoyl-3-hydroxydecanoate (mono-rhamnolipid), and l-rhamnosyl-l-rhamnosyl-3-hydroxydecanoyl-3-hydroxydecanoate (di-rhamnolipid) (13, 14, 44, 46). HAA is synthesized via the RhlA enzyme and is converted to mono-rhamnolipid by the RhlB enzyme (14, 39). Mono-rhamnolipid is converted to di-rhamnolipid by the RhlC enzyme (44). The rlhAB operon and rhlC gene are induced by acyl homoserine lactone-activated RhlR and are thus under quorum-sensing control (39-42, 44). In addition, phosphate limitation and the presence of nitrate have been shown to promote the synthesis of rhamnolipids, while ammonium and high amounts of iron have been shown to repress the production of rhamnolipids (18, 37, 38).

The formation of the mushroom-shaped multicellular structures in P. aeruginosa biofilms under some conditions occurs in a sequential process which involves a nonmotile bacterial subpopulation that forms the mushroom stalks by clonal cell proliferation in certain foci on the substratum and a migrating bacterial subpopulation that forms the mushroom caps via a process which requires type IV pili (26). Type IV pili have been implicated in two kinds of surface-associated motility in P. aeruginosa. One form of surface-associated translocation of P. aeruginosa which requires functional flagella and biosurfactant production, and under some conditions type IV pili, has been termed swarming motility (29, 45, 49). Another form of surface-associated translocation of P. aeruginosa, which requires type IV pili, has been termed twitching motility (20, 34, 48). Presently, it is not known whether the type IV pili-driven motility that plays a role in mushroom cap formation in P. aeruginosa biofilms should be regarded as twitching, swarming, or another kind of surface-associated motility.

Because surface-associated motility and biosurfactant production evidently both have roles in structural biofilm formation by P. aeruginosa, we found it of interest to investigate whether the biosurfactants produced by P. aeruginosa might play a role in structural biofilm development by facilitating surface-associated motility. We present evidence that migration-dependent formation of the cap portion of the mushroom-shaped structures in P. aeruginosa biofilms is facilitated by biosurfactant production. In addition, we present evidence that biosurfactant production is necessary for initial microcolony formation in P. aeruginosa biofilms.


Bacterial strains, plasmids, and growth conditions.

The bacterial strains and plasmids used in this study are listed in Table Table1.1. Escherichia coli strains were cultured in Luria broth (LB) medium at 37°C. P. aeruginosa strains were cultured in LB medium at 37°C during the procedure of genetic manipulation. In motility plate assays, orcinol assays, and for cultivation of biofilms, AB minimal medium supplemented with glucose as indicated was used. AB medium consists of (NH4)2SO4 (15.1 mM), Na2HPO4·2H2O (33.7 mM), KH2PO4 (22.0 mM), NaCl (0.051 M), MgCl2 (1 mM), CaCl2 (0.1 mM), and trace metals (100 μl/liter). The trace metal solution contained CaSO4·2H2O (200 mg/liter), FeSO4·7H2O (200 mg/liter), MnSO4·H2O (20 mg/liter), CuSO4·5H2O (20 mg/liter), ZnSO4·7H2O (20 mg/liter), CoSO4·7H2O (10 mg/liter), NaMoO4·H2O, and H3BO3 (5 mg/liter). Antimicrobial agents were used where appropriate at the following concentrations: for E. coli, ampicillin (Vepidan ApS, Denmark) at 100 μg/ml; for P. aeruginosa, gentamicin sulfate (Biochrome AG, Germany) at 30 μg/ml, streptomycin sulfate (Sigma) at 300 μg/ml, potassium tellurite (Sigma) at 150 μg/ml, and carbenicillin (Sigma) at 200 μg/ml. The P. aeruginosa PAO1 (23) strain from John Matticks' laboratory and an isogenic pilA::Telr derivative (27) were used in this study. The rhlA mutant strain was obtained by transferring the mutated rhlA gene from P. aeruginosa PAO1 rhlA::Gmr (44) (kindly provided by G. A. O'Toole) into P. aeruginosa PAO1, using the transducing phage E79tv2 (36). The P. aeruginosa PAO1 pilA rhlA double mutant strain was derived from the rhlA mutant by allelic displacement of pilA with pilA::Telr using the knockout plasmid pTTN80. The strains were fluorescently tagged at an intergenic neutral chromosomal locus downstream of the glmS gene with ecfp, eyfp, or egfp in mini-Tn7 constructs as described previously (31). Plasmids were transformed into P. aeruginosa strains using electroporation (25 μF, 200 Ω, <5 ms, 2.5 kV).

Strains, plasmids, and primers used in this study

Rhamnolipid assay.

The concentration of rhamnolipids in culture supernatants was determined by the orcinol method as previously described (5, 28, 47), with modifications. Briefly, P. aeruginosa strains were grown at 30°C for 4 days in AB minimal medium supplemented with 10 mM glucose. A 0.5-ml aliquot of culture supernatant was extracted twice with 2 volumes of diethyl ether (high-performance liquid chromatography grade; Sigma). The ether fractions were pooled, evaporated to dryness, and reconstituted in 0.25 ml distilled H2O. A 100-μl aliquot of each sample was diluted 1:10 in orcinol reagent and heated at 80°C for 30 min. Orcinol reagent was prepared immediately prior to use and consisted of 7.5 volumes of 60% (vol/vol) sulfuric acid and 1 volume of 1.6% (wt/vol) orcinol in distilled water. After heating, the samples were allowed to cool at room temperature for 15 min, and absorbance (A421) was measured and compared with rhamnose standards. The concentration of rhamnolipids was determined by the relation that 1.0 mg of rhamnose corresponds to 2.5 mg of rhamnolipid (41).

Twitching motility plate assay.

Twitching motility was assayed on plates composed of AB minimal medium supplemented with 10 mM glucose and solidified with 1.0% agar. The plates were dried overnight at room temperature and point inoculated through the agar surface with 2.5-μl aliquots taken from overnight cultures of P. aeruginosa. Overnight cultures of P. aeruginosa were grown in AB minimal medium, supplemented with 30 mM glucose, at 30°C under vigorous shaking. All plates were incubated at 30°C for 48 h. Twitching motility was visualized by staining with Coomassie brilliant blue R250 (Sigma) as described by Semmler et al. (48). To complement the twitching motility-deficient phenotype of the rhlA mutant, 0.0005% Tween 20 was added to the agar medium where indicated. Twitching motility of P. aeruginosa rhlA containing either pEX1.8 or pEX1.8-rhlAB was examined in the presence of carbenicillin.

Cultivation of biofilms.

Biofilms were cultivated in three-channel flow cells with individual channel dimensions of 1 by 4 by 40 mm and covered with a glass coverslip serving as substratum for biofilm formation. The flow system was assembled and prepared as described elsewhere (50). AB minimal medium supplemented with 0.3 mM glucose as carbon source was used as growth medium. Individual flow chambers were inoculated with 300-μl aliquots taken from overnight cultures of P. aeruginosa, adjusted to an optical density at 500 nm of 0.005. Overnight cultures of P. aeruginosa strains were grown in AB minimal medium supplemented with 30 mM glucose at 30°C under vigorous shaking. P. aeruginosa rhlA strains containing either pEX1.8 or pEX1.8-rhlAB were grown as overnight cultures in the presence of carbenicillin. After inoculation, flow chambers were left without flow for 1 h to allow bacterial attachment. The flow system was incubated at 30°C, and a laminar flow with a mean flow velocity of 0.2 mm s−1 was achieved using a Watson Marlow 205S peristaltic pump. P. aeruginosa rhlA strains harboring either pEX1.8 or pEX1.8-rhlAB were cultivated as biofilms without supplementation of carbenicillin. Loss of plasmid during 4 days of growth in biofilms was not observed as examined by plating of cells derived from biofilms on LB agar medium with and without carbenicillin.

Microscopy and image processing.

Image acquisition was performed by the use of a Zeiss LSM 510 confocal microscope (Carl Zeiss, Jena, Germany) equipped with an argon laser and with detectors and filter sets for simultaneous monitoring of cyan fluorescent protein (CFP) and yellow fluorescent protein (YFP). Spectral imaging of mixed three-color-coded biofilms (CFP, green fluorescent protein [GFP], and YFP) was carried out using the Zeiss META module. Lambda stacks within a spectral range from 453.5 to 549.8 nm were recorded using laser excitation at 458, 488, and 514 nm. Reference emission spectra from single-color-coded biofilms were used for linear unmixing of the mixed color-coded biofilms. Images were obtained using a 40×/1.3 Plan-Neofluar oil objective. Simulated three-dimensional images, shadow projections, and vertical cross-sections were generated using the Imaris software package (Bitplane AG).


In order to examine the effect of biosurfactant production on biofilm development by P. aeruginosa, we constructed isogenic P. aeruginosa strains with and without the capability to produce biosurfactants. We inactivated the rhlA gene in our P. aeruginosa PAO1 strain by the use of transduction with phages that had been propagated on a P. aeruginosa rhlA::Gmr strain. Insertion of the rhlA::Gmr cassette in our PAO1 strain was confirmed by PCR, and deficiency of biosurfactant production was confirmed by a drop-collapsing assay (data not shown). In addition, an orcinol assay showed that the wild-type PAO1 produced 50.9 μg rhamnolipid/ml (n = 3; standard deviation [SD], 7.1) when it was grown in the medium used to cultivate biofilms, compared to 0.58 μg rhamnolipid/ml (n = 3; SD, 1.1) measured in cultures of the rhlA mutant. As biosurfactants have been described to facilitate swarming motility (25), we examined the ability of the PAO1 rhlA mutant and wild type to swarm on semisolid glucose minimal medium and found that the rhlA mutant was deficient in swarming, as expected (data not shown). Subsequently, we tested the ability of the rhlA mutant to perform twitching motility and found that the rate of expansion at the interstitial phase between a plastic surface and agar was significantly reduced for the rhlA mutant compared to the wild type (Fig. (Fig.1A1A and Table Table2),2), suggesting that biosurfactants can also facilitate twitching motility. The P. aeruginosa pilA mutant (which does not produce type IV pili) was included as a nontwitching control (Fig. (Fig.1A).1A). The twitching defect of the rhlA mutant was complemented in P. aeruginosa PAO1 rhlA(pEX1.8-rhlAB), which contains a plasmid-borne rhlAB operon (Fig. (Fig.1B1B and Table Table2).2). To further substantiate that surfactants can facilitate twitching motility, we investigated if the presence of the surfactant Tween 20 could restore twitching motility of the rhlA mutant in the plate assay. In the presence of very low concentrations of Tween 20, the rhlA mutant was able to migrate by twitching motility at the plastic-agar interstitial surface to the same extent as the wild type (with or without Tween 20) (Fig. (Fig.1C1C and Table Table22).

FIG. 1.
Twitching motility plate assays. Glucose minimal medium agar plates were point inoculated with P. aeruginosa PAO1 strains and incubated at 30°C for 48 h. The twitching zones were visualized by staining with Coomassie brilliant blue after incubation. ...
Quantification of twitching motility

We have previously found that when P. aeruginosa was grown in flow chambers with glucose as carbon source, the bacteria differentiated into a nonmotile and a motile subpopulation and formed mushroom-shaped multicellular structures with the nonmotile subpopulation in the stalk portion and the motile subpopulation eventually settling in the cap portion (26). When a biofilm was initiated with a mixture of P. aeruginosa wild type and P. aeruginosa pilA mutant, the nonmotile pilA mutant cells formed mushroom stalks only, whereas the motile wild-type bacteria were able to form mushroom caps on top of the mushroom stalks formed by the pilA mutants (26). To address whether biosurfactants may facilitate cap formation by the motile subpopulation in P. aeruginosa biofilms, we inoculated glucose minimal medium-perfused flow chambers with 1:1 mixtures of CFP-tagged pilA mutant and either YFP-tagged wild type or YFP-tagged rhlA mutant and investigated structural biofilm development in the flow chambers by the use of confocal laser scanning microscopy (CLSM). After 4 days, the wild type had formed cap-shaped structures on top of the stalk-shaped structures formed by the pilA mutant (Fig. (Fig.2A),2A), as has been shown previously by Klausen et al. (26). By day 4 the rhlA mutant also had formed cap-shaped structures on top of the stalk-forming pilA mutant, but the caps formed by the rhlA mutant were significantly smaller than those formed by the wild type (Fig. (Fig.2).2). The defect of the rhlA mutant in cap structure formation could be complemented by introducing the plasmid pEX1.8-rhlAB, expressing the rhlAB operon in trans. The rhlA(pEX1.8-rhlAB) strain formed caps on top of the stalk-forming pilA mutant which were similar to those formed by the wild type, whereas the rhlA(pEX1.8) strain formed caps similar to those formed by the rhlA mutant (Fig. (Fig.2).2). Analysis of 50 randomly chosen mushroom-shaped structures in each of the 4-day-old mixed-strain biofilms showed that the wild type formed caps with an average height of 65.34 μm (SD, 9.80), while the rhlA mutant formed caps with an average height of 31.30 μm (SD, 5.99). The complemented strain rhlA(pEX1.8-rhlAB) formed caps with an average height of 65.71 μm (SD, 10.37), whereas the vector control strain rhlA(pEX1.8) formed caps with an average height of 27.02 μm (SD, 5.28). The experiment therefore suggested that biosurfactants produced by P. aeruginosa have a role in facilitating mushroom cap formation.

FIG. 2.
Confocal laser scanning micrographs of 4-day-old biofilms, initiated with a 1:1 mixture of either P. aeruginosa pilA::Telr Cfp and P. aeruginosa Yfp wild type (A), P. aeruginosa pilA::Telr Cfp and P. aeruginosa PAO1 rhlA::Gmr Yfp (B), P. aeruginosa pilA ...

The experiments indicating that biosurfactants may facilitate cap formation by facilitating bacterial migration also imply that a putative effect of the biosurfactants to a high degree is confined to the biosurfactant-producing subpopulation, as the stalk-forming pilA subpopulation in the experiments described above was capable of synthesizing biosurfactants. If biosurfactant production (i) facilitates colonization of the upper part of P. aeruginosa biofilms by facilitating bacterial migration, and (ii) exerts an effect which to a large extent is confined to the biosurfactant-producing subpopulation, then a mixed biofilm formed by P. aeruginosa pilA, rhlA, and wild-type strains should form multicellular structures with the pilA mutant at the base, the rhlA mutant on top of the pilA mutant, and the wild type on top of the rhlA mutant. To investigate this hypothesis we inoculated flow chambers with a 1:1:1 mixture of CFP-tagged pilA mutant, YFP-tagged rhlA mutant, and GFP-tagged wild type. As shown in Fig. Fig.3,3, in support of the hypothesis, (i) the pilA mutants, which can produce biosurfactants but are unable to migrate, formed microcolonies at the substratum, (ii) the rhlA mutants, which cannot produce biosurfactants and in a twitching motility assay showed reduced ability to migrate, were able to colonize on top of the microcolonies formed by the pilA mutants, and (iii) the wild-type bacteria, which are capable of producing biosurfactants and have no migration deficiency, were able to colonize on top of the rhlA mutants. Because the fluorescence emission peaks of CFP, GFP, and YFP are close to each other, simultaneous detection of these three fluorescent proteins is not possible by the use of conventional CLSM. Therefore, simultaneous detection of the YFP-, GFP-, or CFP-tagged cells in the biofilm was carried out by the use of a special detector system mounted on the microscope (as described in Materials and Methods) which, as apparent in Fig. Fig.3,3, has diminished signal detection compared to conventional CLSM.

FIG. 3.
Confocal laser scanning micrographs of biofilms initiated with a 1:1:1 mixture of P. aeruginosa pilA::Telr Cfp, P. aeruginosa rhlA::Gmr Yfp, and P. aeruginosa PAO1 Gfp wild type. Representative vertical sections acquired at three different locations in ...

Although the effect of the biosurfactants apparently to a high degree is confined to the biosurfactant-producing subpopulation, we could not exclude that biosurfactants produced by the pilA or wild-type subpopulations in the mixed-strain biofilms to some extent could facilitate migration of the rhlA mutant. If biosurfactants were necessary for the motile subpopulation to migrate on the pilA microcolonies, we would expect that the rhlA mutant would not be able to colonize pilA microcolonies which are not capable of providing biosurfactants. To address this, we knocked out the pilA gene in the rhlA mutant by the use of allelic exchange, inoculated a flow cell with a 1:1 mixture of CFP-tagged pilA rhlA double mutant and YFP-tagged rhlA mutant, and investigated biofilm development in glucose minimal medium. We expected that the pilA rhlA double mutant would form mushroom stalks and that we would be able to investigate the ability of the rhlA mutant to form mushroom caps in a biofilm entirely devoid of biosurfactants. Surprisingly, however, microcolonies did not develop in the biofilm containing the mixture of the pilA rhlA double mutant and the rhlA mutant. Instead, the bacteria formed a flat biofilm, as shown in Fig. Fig.4,4, which depicts a CLSM micrograph acquired in a 4-day-old pilA rhlA-rhlA mixed biofilm.

FIG. 4.
Confocal laser scanning micrograph of a 4-day-old biofilm, initiated with a 1:1 mixture of P. aeruginosa rhlA::Gmr Yfp and P. aeruginosa pilA::Telr rhlA::Gmr Cfp. The image shows a top-down shadow projection (230 μm by 230 μm) with two ...

Because our experiments with mixed-strain P. aeruginosa biofilms surprisingly had indicated that biosurfactants, in addition to their role in the formation of mature biofilm structures, might also play a role in the formation of the initial microcolonies, we found it of interest to examine the role of biosurfactant production on biofilm development in more simple model systems consisting of mono-strain P. aeruginosa biofilms. While the wild type and the pilA mutant after 4 days of development had formed a biofilm with mushroom-shaped structures (Fig. (Fig.5A)5A) and irregular protruding structures (Fig. (Fig.5C),5C), respectively, the rhlA mutant and the pilA rhlA double mutant both formed flat biofilms (Fig. 5B and D), supporting the hypothesis that biosurfactants are necessary for initial microcolony formation.

FIG. 5.
Confocal laser scanning micrographs of 4-day-old biofilms formed by P. aeruginosa PAO1 Yfp wild type (A), P. aeruginosa rhlA::Gmr Yfp (B), P. aeruginosa pilA::Telr Cfp (C), and P. aeruginosa pilA::Telr rhlA::Gmr Cfp (D). The images show top-down shadow ...

To examine further the role of biosurfactants in initial microcolony formation, we examined young P. aeruginosa biofilms. After 16 h of development in glucose minimal medium-perfused flow chambers, the wild type had formed small microcolonies (Fig. (Fig.6A),6A), whereas the rhlA mutant had formed a flat thin biofilm (Fig. (Fig.6B).6B). When the rhlA mutant and the wild type were present in the flow chamber in a 1:1 mixture, the rhlA mutant was able to form initial microcolonies of the same size as those formed by the wild type (Fig. (Fig.6C),6C), suggesting that biosurfactant produced by the wild type enabled microcolony formation by the rhlA mutant.

FIG. 6.
Confocal laser scanning micrographs of 16-h-old biofilms formed by P. aeruginosa PAO1 Cfp wild type (A), P. aeruginosa rhlA::Gmr Yfp (B), and a 1:1 mixture of P. aeruginosa PAO1 Cfp wild type and P. aeruginosa rhlA::Gmr Yfp (C). The images show top-down ...


Because previous studies had indicated that surface-associated motility and biosurfactant production both play roles in structure formation in P. aeruginosa biofilms (10, 26), we found it of interest to investigate if the role of biosurfactants in structural P. aeruginosa biofilm formation can be attributed to facilitation of bacterial migration. As discussed below, our results indicate that biosurfactants produced by P. aeruginosa indeed do play roles in migration-mediated structure formation in the later stages of biofilm development and, in addition, have an impact on microcolony formation in the early phase of P. aeruginosa biofilm development.

Initial studies to determine the capability of an rhlA mutant to migrate in conventional motility plate assays suggested that biosurfactants promote twitching motility of P. aeruginosa. The P. aeruginosa rhlA mutant used in the present study and other P. aeruginosa rhlA mutants of different PAO1 sublines were found to be impaired in twitching motility compared to their isogenic wild type (Fig. (Fig.1A1A and data not shown). The defect of the rhlA mutant strains in migration in the twitch plate assay could in each case be complemented by either introducing the biosurfactant biosynthesis genes rhlAB in trans or by providing the surfactant Tween 20 (Fig. 1B and C and data not shown). The observed defect of P. aeruginosa rhlA mutants in migration in twitch plate assays was to some extent surprising, since so far biosurfactants have only been reported to facilitate motility by P. aeruginosa in swarm plate assays (14, 29). However, it appears that the ability of biosurfactants to reduce surface tension and act as wetting agents may facilitate different kinds of surface-associated translocation of P. aeruginosa. In addition, it has been reported that biosurfactants may affect the composition of lipopolysaccharide (LPS) on the outer membrane of P. aeruginosa (2) and that a change in the LPS composition of P. aeruginosa may cause a significant reduction in the capability of the bacterium to migrate in both twitch plate and swarm plate assays (1). An effect of changes in cell surface LPS composition on migration in motility plate assays has been observed for several species, including Salmonella enterica, Proteus mirabilis, and Myxococcus xanthus (6, 17, 35, 53, 55). In the case of S. enterica, it was shown that the defect in motility due to changes in the LPS composition could be restored by providing a surfactant (53).

To investigate the impact of biosurfactants on migration-mediated structural biofilm development, we performed experiments with mixed-strain color-coded biofilms. Our results provide evidence that biosurfactants produced by P. aeruginosa indeed facilitate bacterial migration and colonization of the upper part of the biofilm. The rhlA mutant developed mushroom caps of reduced size on top of mushroom stalks formed by the pilA mutant, compared to the mushroom caps developed by the wild type in an analogous setup. In addition, experiments with three-strain color-coded biofilms suggested that, unlike the wild-type strain, the motility-impaired rhlA mutant was not able to colonize the top of the mushroom-shaped structures. These experiments also indicated that the possible effects of biosurfactant on cellular migration and structure formation are largely confined to the subpopulations that produce the biosurfactants. It is likely that, due to their inherent amphipathic characteristics, the biosurfactants will adhere directly to the cell surfaces of the producing cells or to a cell surface in close proximity and exert their effect on cellular migration, leading to the spatial distribution of distinct subpopulations within the multicellular structures. Because biosurfactant production evidently may facilitate both swarming and twitching motility, the involvement of biosurfactants in facilitating migration-dependent structural development in P. aeruginosa biofilms does not provide information regarding the type of surface-associated motility occurring.

In agreement with our observations, Davey et al. (10) reported that a P. aeruginosa rhlA mutant formed a flat biofilm. Contrary to our suggestion, however, Davey et al. (10) suggested that the biosurfactants are not required for the formation of initial microcolonies but that they participate in the maintenance of the channels between the microcolonies once they are formed. When we used the same P. aeruginosa rhlA mutant as was used by Davey and colleagues we got essentially the same results as reported by them. That is, this P. aeruginosa rhlA strain indeed formed microcolonies and the channels between the microcolonies were subsequently colonized (data not shown). However, when we moved the rhlA mutation via phage transduction to a number of different PAO1 sublines, the newly constructed rhlA mutants were all deficient in microcolony formation as reported here. This suggests that the rhlA mutant used by Davey et al. (10) carries additional mutations that enable it to form microcolonies in the absence of biosurfactants. However, differences in media and experimental setup may also be the cause of these seemingly divergent results.

A role of P. aeruginosa biosurfactants in initial microcolony formation appears to be contrary to the established roles of biosurfactants in maintaining channels between microcolonies (10) and in dispersal of biofilm (5, 24, 47). However, in support of the proposed role of P. aeruginosa biosurfactant in initial microcolony formation, it has been shown that low concentrations of rhamnolipid enhance the cell surface hydrophobicity of P. aeruginosa by causing a release of LPS from the cell surface (2, 56). An increase in cell surface hydrophobicity could increase the adhesiveness of the bacteria to a level which is critical for initial microcolony formation in biofilms. In support of this suggestion, Herman et al. (22) showed that addition of low concentrations of rhamnolipid induced the formation of multicellular aggregates in P. aeruginosa suspensions. It is possible that the bacteria in wild-type P. aeruginosa biofilms produce low amounts of biosurfactants, sufficient to facilitate microcolony formation, in the early phase of biofilm development when the cell concentration is still not high enough to constitute a quorum. When the cell concentration reaches higher levels, the production of high concentrations of P. aeruginosa biosurfactants is expected to be induced via quorum sensing, and these high concentrations of P. aeruginosa biosurfactants might have the proposed effects on cellular migration, channel maintenance, and biofilm dispersal. At present we cannot explain why the effect of biosurfactants on structure formation in developed biofilms seemed to be confined largely to biosurfactant-producing subpopulations, whereas biosurfactant production by the wild type in young wild-type-rhlA mixed biofilms apparently could facilitate microcolony formation of both the wild type and the rhlA mutant. However, this might be related to differences in the amount and composition of biosurfactants required in the different processes. Caiazza and coworkers recently described that the various biosurfactant compounds produced by P. aeruginosa have different effects on cell surface hydrophobicity and cellular migration (7).

Due to the occurrence of nutrient and oxygen gradients in biofilms, the top of the mushroom-shaped structures is believed to be a favored location, because the cells present there will receive more substrate from the bulk liquid than the cells situated within the multicellular structures or close to the substratum (12, 16, 30, 43, 54). It appears that only cells which are fully capable of type IV pilus-driven migration succeed in reaching the favorable top of the mushroom-shaped structures. Experiments to investigate a possible role of chemotaxis in coordinating the cellular migration involved in cap formation are under way in our laboratory.

In conclusion, the present study together with the studies of others suggest that P. aeruginosa biosurfactants have multiple roles in P. aeruginosa biofilm development: (i) they are necessary for initial microcolony formation, (ii) they facilitate surface-associated bacterial migration and thereby the formation of mushroom-shaped structures, (iii) they prevent colonization of the channels between the mushroom-shaped structures (10), and (iv) they play a role in biofilm dispersion (5, 47).


This work was supported by grants from the Danish Research Agency.

We thank G. A. O'Toole for providing a P. aeruginosa rhlA strain and for critical reading of the manuscript. We are grateful to P. K. Singh for providing the plasmids pEX1.8 and pEX1.8-rhlAB.


[down-pointing small open triangle]Published ahead of print on 12 January 2007.


1. Abeyrathne, P. D., C. Daniels, K. K. Poon, M. J. Matewish, and J. S. Lam. 2005. Functional characterization of WaaL, a ligase associated with linking O-antigen polysaccharide to the core of Pseudomonas aeruginosa lipopolysaccharide. J. Bacteriol. 187:3002-3012. [PMC free article] [PubMed]
2. 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. [PMC free article] [PubMed]
3. Banin, E., K. M. Brady, and E. P. Greenberg. 2006. Chelator-induced dispersal and killing of Pseudomonas aeruginosa cells in a biofilm. Appl. Environ. Microbiol. 72:2064-2069. [PMC free article] [PubMed]
4. Bao, Y., D. P. Lies, H. Fu, and G. P. Roberts. 1991. An improved Tn7-based system for the single-copy insertion of cloned genes into chromosomes of gram-negative bacteria. Gene 109:167-168. [PubMed]
5. Boles, B. R., M. Thoendel, and P. K. Singh. 2005. Rhamnolipids mediate detachment of Pseudomonas aeruginosa from biofilms. Mol. Microbiol. 57:1210-1223. [PubMed]
6. Bowden, M. G., and H. B. Kaplan. 1998. The Myxococcus xanthus lipopolysaccharide O-antigen is required for social motility and multicellular development. Mol. Microbiol. 30:275-284. [PubMed]
7. Caiazza, N. C., R. M. Shanks, and G. A. O'Toole. 2005. Rhamnolipids modulate swarming motility patterns of Pseudomonas aeruginosa. J. Bacteriol. 187:7351-7361. [PMC free article] [PubMed]
8. Costerton, J. W., P. S. Stewart, and E. P. Greenberg. 1999. Bacterial biofilms: a common cause of persistent infections. Science 284:1318-1322. [PubMed]
9. Costerton, J. W., Z. Lewandowski, D. E. Caldwell, D. R. Korber, and H. M. Lappin-Scott. 1995. Microbial biofilms. Annu. Rev. Microbiol. 49:711-745. [PubMed]
10. 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. [PMC free article] [PubMed]
11. De Beer, D., R. Srinivasan, and P. S. Stewart. 1994. Direct measurement of chlorine penetration into biofilms during disinfection. Appl. Environ. Microbiol. 60:4339-4344. [PMC free article] [PubMed]
12. De Beer, D., P. Stoodley, F. Roe, and Z. Lewandowski. 1994. Effects of biofilm structures on oxygen distribution and mass transport. Biotechnol. Bioeng. 43:1131-1138. [PubMed]
13. Deziel, E., F. Lepine, D. Dennie, D. Boismenu, O. A. Mamer, and R. Villemur. 1999. Liquid chromatography/mass spectrometry analysis of mixtures of rhamnolipids produced by Pseudomonas aeruginosa strain 57RP grown on mannitol or naphthalene. Biochim. Biophys. Acta 1440:244-252. [PubMed]
14. 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-hydroxy-alkanoyloxy)alkanoic acids (HAAs), the precursors of rhamnolipids. Microbiology 149:2005-2013. [PubMed]
15. Donlan, R. M., and J. W. Costerton. 2002. Biofilms: survival mechanisms of clinically relevant microorganisms. Clin. Microbiol. Rev. 15:167-193. [PMC free article] [PubMed]
16. Eberl, H. J., D. F. Parker, and M. C. M. van Loosdrecht. 2001. A new deterministic spatio/temporal continuum model for biofilm development. J. Theor. Med. 3:161-175.
17. Gue, M., V. Dupont, A. Dufour, and O. Sire. 2001. Bacterial swarming: a biochemical time-resolved FTIR-ATR study of Proteus mirabilis swarm-cell differentiation. Biochemistry 40:11938-11945. [PubMed]
18. Guerra-Santos, L., O. Kappeli, and A. Fiechter. 1984. Pseudomonas aeruginosa biosurfactant production in continuous culture with glucose as carbon source. Appl. Environ. Microbiol. 48:301-305. [PMC free article] [PubMed]
19. Haagensen, J. A., M. Klausen, R. K. Ernst, S. I. Miller, A. Folkesson, T. Tolker-Nielsen, and S. Molin. 2007. Differentiation and distribution of colistin- and sodium dodecyl sulfate-tolerant cells in Pseudomonas aeruginosa biofilms. J. Bacteriol. 189:28-37. [PMC free article] [PubMed]
20. Henrichsen, J. 1972. Bacterial surface translocation: a survey and a classification. Bacteriol. Rev. 36:478-503. [PMC free article] [PubMed]
21. Hentzer, M., H. Wu, J. B. Andersen, K. Riedel, T. B. Rasmussen, N. Bagge, N. Kumar, M. A. Schembri, Z. Song, P. Kristoffersen, M. Manefield, J. W. Costerton, S. Molin, L. Eberl, P. Steinberg, S. Kjelleberg, N. Hoiby, and M. Givskov. 2003. Attenuation of Pseudomonas aeruginosa virulence by quorum sensing inhibitors. EMBO J. 22:3803-3815. [PMC free article] [PubMed]
22. Herman, D. C., Y. Zhang, and R. M. Miller. 1997. Rhamnolipid (biosurfactant) effects on cell aggregation and biodegradation of residual hexadecane under saturated flow conditions. Appl. Environ. Microbiol. 63:3622-3627. [PMC free article] [PubMed]
23. Holloway, B. W., and A. F. Morgan. 1986. Genome organization in Pseudomonas. Annu. Rev. Microbiol. 40:79-105. [PubMed]
24. Irie, Y., G. A. O'Toole, and M. H. Yuk. 2005. Pseudomonas aeruginosa rhamnolipids disperse Bordetella bronchiseptica biofilms. FEMS Microbiol. Lett. 250:237-243. [PubMed]
25. Kessler, B., V. de Lorenzo, and K. N. Timmis. 1992. A general system to integrate lacZ fusions into the chromosomes of gram-negative eubacteria: regulation of the Pm promoter of the TOL plasmid studied with all controlling elements in monocopy. Mol. Gen. Genet. 233:293-301. [PubMed]
26. Klausen, M., A. Aaes-Jørgensen, 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. [PubMed]
27. 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. [PubMed]
28. Koch, A. K., O. Kappeli, A Fiechter, and J Reiser. 1991. Hydrocarbon assimilation and biosurfactant production in Pseudomonas aeruginosa mutants. J. Bacteriol. 173:4212-4219. [PMC free article] [PubMed]
29. Köhler, 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. [PMC free article] [PubMed]
30. Kreft, J. U., C. Picioreanu, J. W. Wimpenny, and M. C. van Loosdrecht. 2001. Individual-based modelling of biofilms. Microbiology 147:2897-2912. [PubMed]
31. Lambertsen, L., C. Sternberg, and S. Molin. 2004. Mini-Tn7 transposons for site-specific tagging of bacteria with fluorescent proteins. Environ. Microbiol. 6:726-732. [PubMed]
32. Lawrence, J. R., D. R. Korber, B. D. Hoyle, J. W. Costerton, and D. E. Caldwell. 1991. Optical sectioning of microbial biofilms. J. Bacteriol. 173:6558-6567. [PMC free article] [PubMed]
33. Lequette, Y., and E. P. Greenberg. 2005. Timing and localization of rhamnolipid synthesis gene expression in Pseudomonas aeruginosa biofilms. J. Bacteriol. 187:37-44. [PMC free article] [PubMed]
34. Mattick, J. S. 2002. Type IV pili and twitching motility. Annu. Rev. Microbiol. 56:289-314. [PubMed]
35. McCoy, A. J., H. Liu, T. J. Falla, and J. S. Gunn. 2001. Identification of Proteus mirabilis mutants with increased sensitivity to antimicrobial peptides. Antimicrob. Agents Chemother. 45:2030-2037. [PMC free article] [PubMed]
36. Morgan, A. F. 1979. Transduction of Pseudomonas aeruginosa with a mutant of bacteriophage E79tv2. J. Bacteriol. 139:137-140. [PMC free article] [PubMed]
37. Mulligan, C. N., and B. F. Gibbs. 1989. Correlation of nitrogen metabolism with biosurfactant production by Pseudomonas aeruginosa. Appl. Environ. Microbiol. 55:3016-3019. [PMC free article] [PubMed]
38. Mulligan, C. N., G. Mahmourides, and B. F. Gibbs. 1989. The influence of phosphate metabolism on biosurfactant production by Pseudomonas aeruginosa. J. Biotechnol. 12:199-210.
39. Ochsner, U. A., A. Fiechter, and J. Reiser. 1994. Isolation, characterization, and expression in Escherichia coli of the Pseudomonas aeruginosa rhlAB genes encoding a rhamnosyltransferase involved in rhamnolipid biosurfactant synthesis. J. Biol. Chem. 269:19787-19795. [PubMed]
40. Ochsner, U. A., and J. Reiser. 1995. Autoinducer-mediated regulation of rhamnolipid biosurfactant synthesis in Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. USA 92:6424-6428. [PMC free article] [PubMed]
41. Pearson, J. P., E. C. Pesci, and B. H. Iglewski. 1997. Roles of Pseudomonas aeruginosa las and rhl quorum-sensing systems in control of elastase and rhamnolipid biosynthesis genes. J. Bacteriol. 179:5756-5767. [PMC free article] [PubMed]
42. Pesci, E. C., J. P. Pearson, P. C. Seed, and B. H. Iglewski. 1997. Regulation of las and rhl quorum sensing in Pseudomonas aeruginosa. J. Bacteriol. 179:3127-3132. [PMC free article] [PubMed]
43. Picioreanu, C., M. C. M. van Loosdrecht, and J. J. Heijnen. 1998. Mathematical modeling of biofilm structure with a hybrid differential-discrete cellular automaton approach. Biotechnol. Bioeng. 58:101-116. [PubMed]
44. 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. [PubMed]
45. 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 25:4885-4890. [PMC free article] [PubMed]
46. Rendell, N. B., G. W. Taylor, M. Somerville, H. Todd, R. Wilson, and P. J. Cole. 1990. Characterisation of Pseudomonas rhamnolipids. Biochim. Biophys. Acta 16:189-193. [PubMed]
47. Schooling, S. R., U. K. Charaf, D. G. Allison, and P. Gilbert. 2004. A role for rhamnolipid in biofilm dispersion. Biofilms 1:91-99.
48. Semmler, A. B., C. B. Whitchurch, and J. S. Mattick. 1999. A re-examination of twitching motility in Pseudomonas aeruginosa. Microbiology 145:2863-2873. [PubMed]
49. Sharma, M., and S. K. Anand. 2002. Swarming: a coordinated bacterial activity. Curr. Sci. 83:707-715.
50. Sternberg, C., and T. Tolker-Nielsen. 2005. Growing and analyzing biofilms in flow cells, p. 1B.2.1-1B.2. 15. In R. Coico, T. Kowalik, J. Quarles, B. Stevenson, and R. Taylor (ed.), Current protocols in microbiology. John Wiley & Sons, Inc., New York, N.Y.
51. Stewart, P. S., and J. W. Costerton. 2001. Antibiotic resistance of bacteria in biofilms. Lancet 14:135-138. [PubMed]
52. Stoodley, P., D. De Beer, and Z. Lewandowski. 1994. Liquid flow in biofilm systems. Appl. Environ. Microbiol. 60:2711-2716. [PMC free article] [PubMed]
53. Toguchi, A., M. Siano, M. Burkart, and R. M. Harshey. 2000. Genetics of swarming motility in Salmonella enterica serovar Typhimurium: critical role for lipopolysaccharide. J. Bacteriol. 182:6308-6321. [PMC free article] [PubMed]
54. Wimpenny, J. W. T., and R. Colasanti. 1997. A unifying hypothesis for the structure of microbial biofilms based on cellular automated models. FEMS Microbiol. Ecol. 22:1-16.
55. Yang, Z., D. Guo, M. G. Bowden, H. Sun, L. Tong, Z. Li, A. E. Brown, H. B. Kaplan, and W. Shi. 2000. The Myxococcus xanthus wbgB gene encodes a glycosyltransferase homologue required for lipopolysaccharide O-antigen biosynthesis. Arch. Microbiol. 174:399-405. [PubMed]
56. 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. [PMC free article] [PubMed]

Articles from Journal of Bacteriology are provided here courtesy of American Society for Microbiology (ASM)
PubReader format: click here to try


Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...


Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...