INTRODUCTION

Top Abstract Introduction Materials and Methods Results Discussion References

Pseudomonas aeruginosa is a gram-negative bacterium found in almost every ecological niche, including soil, water, and plants. It is also an important opportunistic pathogen of humans, primarily infecting immunocompromised patients (17). Recent reports indicate that environmental and clinical P. aeruginosa strains are functionally equivalent and taxonomically indistinguishable (14). The success of P. aeruginosa in various environments is attributed to its broad metabolic versatility and its elaboration of many cell-associated and secreted virulence/survival factors (47).

Among the cell surface structures of P. aeruginosa, the polar flagellum is responsible for a mode of motility in aqueous environment called swimming. As for most other motile bacteria, direction of movement is biased by chemotactic responses to chemical stimuli (6, 31, 46). Flagella also mediate a mode of social motility known as swarming, recently described for the first time in P. aeruginosa (39). Other cell surface structures acting as virulence/survival factors are type IV pili. These polar fimbriae are presumably the principal adhesins, mediating the adherence to eukaryotic cell surfaces (18) and probably to abiotic surfaces as well (41). They are also responsible for the flagellum-independent mode of surface translocation called twitching motility (9, 41, 48).

Bacteria in natural habitats usually grow as biofilms, organized communities of cells embedded in an extracellular polysaccharide matrix and attached to a surface (5). In recent years, much has been learned about how cells initiate biofilm formation (43, 49). Escherichia coli mutants defective in biofilm formation were found either to lack the ability to produce type 1 pili or to be nonmotile (37). Similarly, flagellar motility and type IV pilus-based twitching motility have been shown to be required for the initial attachment and development of a biofilm by P. aeruginosa (34).

Biofilm bacteria display particular phenotypes that distinguish them from their freely growing counterparts (5, 49). The differential expression of a large number of genes is known to occur in the initial steps of biofilm formation (5, 38), such as the upregulation of exopolysaccharide synthesis following bacterial adhesion to a surface (10, 38). However, a regulatory system controlling the conversion to the biofilm phenotype has not been described.

A strategy that bacteria use to rapidly adapt and survive when environmental conditions change is to create a phenotypically diverse population by a mechanism called phase variation, that is, the high-frequency and reversible switching of phenotypic traits (13). In gram-negative bacteria, the expression of a number of cell surface structures and outer membrane proteins, especially those linked to adhesion, aggregation, and colonial morphology, is known to be regulated by phase-variable mechanisms (reviewed in reference 22).

As a part of a research project aimed at understanding the physiological mechanisms used by P. aeruginosa to access and catabolize hydrophobic, poorly bioavailable substrates such petroleum hydrocarbons (11), we have observed the spontaneous emergence of alternate phenotypic forms growing as small, rough colonies when P. aeruginosa 57RP was cultivated as a biofilm or in static liquid cultures (E. Déziel, Y. Comeau, and R. Villemur, Abstr. 99th Gen. Meet. Am. Soc. Microbiol., abstr. K-57, 1999). Since colonial morphology reflects the differential expression of components on cell surfaces within the colony, we hypothesized that the adherence and/or motility behavior of this small colony phenotype might be altered.

In this report, we describe the isolation and characterization of phenotypic variants of P. aeruginosa 57RP. In contrast with the large-colony (L) variant (parent form), the small-colony (S) variants produced abundant polar fimbriae, displayed reduced flagellar (chemotactic) and twitching motilities, and rapidly initiated the formation of strongly adherent biofilms. Under appropriate growth conditions, cells of each phenotype switched to the other phenotype at a fairly high frequency, suggesting that these S variants resulted from a reversible phase variation phenomenon and were selectively enriched when P. aeruginosa was grown as a biofilm or in static liquid cultures.

	MATERIALS AND METHODS

Top Abstract Introduction Materials and Methods Results Discussion References

Bacterial strains and culture media. P. aeruginosa 57RP was originally isolated from a hydrocarbon-contaminated soil (11). Bacteria were routinely subcultured on tryptic soy agar (TSA) plates from frozen stocks, and overnight cultures in Luria-Bertani broth (LB) at 37°C and 250 rpm were used to prepare inocula, unless stated otherwise.

Evaluation of cell surface hydrophobicity and cell adherence. (i) MATH. Estimation of microbial cell surface hydrophobicity was performed with a microbial adhesion to hydrocarbon (MATH) test (40). Cells from overnight cultures were washed twice and resuspended in 25 mM phosphate-buffered saline (PBS) to an optical density at 600 nm of approximately 0.6. Then 1.5 ml of this suspension was mixed with various volumes of hexadecane (ranging from 0 to 700 µl) in 16- by 125-mm test tubes and vortexed for 30 s. After 30 min of equilibration, we measured the loss in absorbance of the aqueous phase relative to that of the initial cell suspension and estimated hydrophobicity by calculating the percentage of cells adhering to hexadecane.

(ii) MATS. To estimate the adhesion potential of the cells (microbial adhesion to silica sand [MATS]), the MATH test was modified by replacing the hexadecane with different amounts (0 to 900 mg) of fine granular silica sand.

Motility and chemotaxis assays. (i) Swimming. Tryptone swim plates (1% tryptone, 0.5% NaCl, 0.3% agar) were inoculated with a sterile toothpick and incubated for 16 h at 25°C. Motility was then assessed qualitatively by examining the circular turbid zone formed by the bacterial cells migrating away from the point of inoculation.

(ii) Swarming. Swarm plates were composed of 0.5% Bacto Agar and 8 g of nutrient broth/liter (both from Difco, Detroit, Mich.), supplemented with 5 g of dextrose/liter, and dried overnight at room temperature (39). Cells were point inoculated with a sterile toothpick, and the plates were incubated at 30°C for 24 h.

(iii) Twitching. Cells were stab inoculated with a toothpick through a thin (approximately 3-mm) LB agar layer (1% agar) to the bottom of the petri dish. After incubation for 24 to 48 h at 30°C, a hazy zone of growth at the interface between the agar and the polystyrene surface was observed (7). The ability of bacteria to strongly adhere and form a biofilm on the polystyrene surface was then examined by removing the agar, washing unattached cells with a stream of tap water, and staining the attached cells with crystal violet (1% [wt/vol] solution).

(iv) Flagellar chemotaxis. The chemotactic response was quantified by a slightly modified version of the capillary assay of Mazumder et al. (32). A 1-ml tuberculin syringe with a disposable 25-gauge needle (Terumo Medical Corp., Elkton, Md.) was filled with 100 µl of Bushnell-Haas (BH) mineral salts medium (Difco) containing 0.1% tryptone as a chemoattractant. Cells were grown in LB at 37°C to the logarithmic phase, washed, and resuspended in BH. A 100-µl sample of this bacterial suspension was drawn into a 200-µl pipette tip. The syringe was then inserted and tightly fit into the tip with 3 mm of the needle inserted into the cell suspension. Control capillaries containing only BH were performed with each assay. Duplicate apparatus were incubated at 37°C for 45 min, and the content of the syringe was then diluted in 25 mM PBS and plated onto TSA plates for cell enumeration.

Biofilm formation assay with polystyrene culture tubes. The biofilm formation protocol was adapted from that of O'Toole and Kolter (35). Polystyrene 12- by 75-mm tubes containing 0.5 ml of BDT medium (BH mineral salts medium supplemented with 0.2% dextrose and 0.5% tryptone) were inoculated from overnight LB cultures and incubated at 32°C without agitation. At regular time intervals, triplicate tubes were rinsed thoroughly with water, and a 1% solution of crystal violet was added to stain the attached cells. After 10 to 15 min of incubation at room temperature, the tubes were rinsed with water, and the biomass of attached cells (biofilm) was quantified by solubilization of the dye in 2 ml of 95% ethanol. The absorbance was measured at 600 nm with a spectrophotometer.

Sensitivity to oxidative stress. The disk assay of Hassett et al. (21) was used to test the sensitivity of cells to oxidative stress. Briefly, 100-µl aliquots from cultures in mid-log or stationary phases of growth were uniformly spread on TSA and medium A (25) plates containing 2% agar. Sterile Whatman no. 1 filter paper disks (7-mm diameter) impregnated with 10 µl of 30% H₂O₂ were placed in triplicate on each plate. The diameter of the zone of growth inhibition around each disk was measured after 5 h of incubation at 37°C.

Production of exoproducts. (i) Pyocyanin. Bacteria were grown for 30 h at 37°C and 250 rpm in 2 ml of medium A, which promotes pyocyanin production, and the relative amount of pyocyanin in culture supernatant was measured spectrophotometrically at 695 nm.

(ii) Pyoverdine. Cells were cultivated at 37°C and 250 rpm for 16 h in 2 ml of medium B (25). The relative concentration of pyoverdine was quantified in the supernatants by measurement of the fluorescence at 460 nm after excitation at 400 nm with a Spectramax Gemini microplate spectrofluorometer (Molecular Devices Corp., Sunnyvale, Calif.).

(iii) Total proteases and elastase. Elastase (LasB protease) activity was determined in liquid cultures by the elastin-Congo red (ECR) hydrolysis assay as described by Pearson et al. (36).

(iv) Determination of alginate production. Cells were cultivated in LB supplemented with 0.2% glycerol for 92 h at 37°C and 250 rpm. The cultures were then centrifuged at 8,000 × g for 5 min, and the alginate contained in the supernatants was precipitated at 70°C for 16 h with 3 volumes of 95% ethanol. The precipitate was recovered by centrifugation at 18,000 × g for 15 min and resuspended in water. Alginate was quantified by assaying uronic acids with the borate-carbazole method (26), with D-mannuronate lactone (Sigma Chemical Co., St. Louis, Mo.) as a standard. Values were normalized to cell growth with total cellular protein concentrations. Proteins were solubilized in 0.1 N NaOH at 70°C for 30 min and analyzed by the method of Bradford (Bio-Rad Laboratories, Hercules, Calif.), with bovine serum albumin as a standard.

(v) Rhamnolipids. Cultures were conducted at 30°C in agitated glass test tubes containing 1 ml of SW1/10F mineral salts medium with 2% mannitol (11, 12). Total production of all isomers of rhamnolipid biosurfactants was estimated in the supernatant by extraction and hydrolysis followed by quantification of rhamnose with the orcinol assay (4). The concentration of rhamnolipids was determined considering that 1 mg of rhamnose corresponds to 2.25 mg of rhamnolipids (12). Because of the highly clumping behavior of the S variants, the whole culture was evaluated for total protein content and correlated to rhamnolipid production.

Electron microscopy. A drop of water was deposited on the edge of a colony from an overnight-grown LB agar plate. Cells were allowed to become suspended for about 1 min; then a Formvar-coated copper grid was floated on the drop for about 45 s, rinsed in a drop of water, and stained for 15 s with a 2% aqueous solution of phosphotungstic acid. Samples were examined with a Hitachi H-7100 transmission electron microscope.

	RESULTS

Top Abstract Introduction Materials and Methods Results Discussion References

Emergence of phenotypic variants of P. aeruginosa 57RP correlates with biofilm formation. The wild-type P. aeruginosa 57RP parent strain usually produces large (~16-mm diameter after 2 days at 30°C), flat colonies with an irregular, finely mottled periphery on LB plates (Fig. 1A). We had previously observed that when this strain was cultivated in liquid medium with hexadecane as the substrate, there was a lag phase (approximately 5 to 10 days) before significant growth occurred. Interestingly, at the onset of the exponential growth phase, we noticed the formation of a biofilm on the surface of hexadecane droplets, and this was correlated with the appearance of small, dry-looking colonies on agar plates. A progression from the wild-type form to the small rough colony phenotype with transient appearance of intermediate size and roughness was also noticed during the cultivation period on hexadecane (Déziel et al., Abstr. 99th Gen. Meet. Am. Soc. Microbiol.).

View larger version (103K): [in this window] [in a new window]   FIG. 1.   Visual differences between growth phenotypes. (A to C) Colonies of L variant (A), S1 variant (B; the arrow indicates the emergence of a L-type revertant sector emerging from the side of a colony), and S2 variant (C) on LB agar plates incubated at 30°C. (D) Overnight growth in broth medium with shaking. The L and S1rev variants grew homogeneously dispersed in the medium, whereas the S1 and S2 variants preferred the interface and the glass surface. S1rev is an L variant resulting from the reversion of an S1 variant.

View larger version (19K): [in this window] [in a new window]   FIG. 2.   Kinetics of biofilm formation. L (), S1 (), and S2 () variants were cultivated in polystyrene tubes at 32°C without agitation. At the indicated time intervals, triplicate tubes were rinsed and stained with crystal violet. The amount of stained cells was then quantified by spectrophotometry (A600) after solubilization of the dye in ethanol.

S variants can revert to the parent L variant phenotype. Outside a selective environment, the S1 variant reverted to the L phenotype at a relatively high frequency. Transfer plating of an S1 colony on TSA plates consistently resulted in the emergence of L-type revertants forming sectors emerging from the colonies (arrow in Fig. 1B). For example, after 1 week of incubation at room temperature, most colonies on a plate displayed outgrowth of L variant cells. The same phenomenon also occurred with the S2 variant but at a much lower frequency, with only occasional revertants appearing on plates. Because of the differences in growth behavior and environmental niche preferences between the L and S variants, determination of switching rates is not readily possible.

S variants demonstrate increased cell surface hydrophobicity and adhesivity. The aggregating and adherent behavior of the S variants suggested that their cell surface hydrophobicity was higher than that of the L variant. According to the standard assay used to quantitatively evaluate cell surface hydrophobicity (MATH test), the cell surface of S1 was much more hydrophobic than the surface of the parent L variant (Fig. 3A). However, this variant was also more adhesive to silica sand, a hydrophilic substratum (Fig. 3B), indicating that the S variants are mainly characterized by their adherence. It was not possible to perform these assays with S2 because of the highly clumping behavior of this variant.

View larger version (18K): [in this window] [in a new window]   FIG. 3.   Evaluation of cell surface hydrophobicity and adhesion potential by the MATH (A) and MATS (B) tests, respectively, in L () and S1 () variants. Various amounts of hexadecane (MATH) or silica sand (MATS) were mixed with a washed cell suspension in PBS, and the optical densities at 600 nm before and after were compared. Values for the MATH test are means ± standard deviations of duplicates.

S variants are deficient in swimming, swarming and twitching motilities. The reduced diameter of S variants colonies suggested that they were impaired in motility and/or chemotaxis. When the S variants were cultivated on soft agar plates, their zones of swimming were smaller than that for the L variant with S2 producing slightly larger zones than S1 (Fig. 4A). There was some dispersion of cells from the point of inoculation but without the formation of concentric chemotactic rings, suggesting that the S variants are not completely defective in motility but may be impaired in chemotaxis. Microscopic examination showed that the S variants were motile, but many cells exhibited a tumbling behavior and random movement and lacked the directional swimming typical of the L phenotype. On swarm agar plates, the S variants remained near the point of inoculation and did not form the expanding and irregular branching pattern which is characteristic of swarming motility in P. aeruginosa, as recently described by Rashid and Kornberg (39) (Fig. 4B).

View larger version (100K): [in this window] [in a new window]   FIG. 4.   Differences in motility phenotypes of L and S variants. (A) Swimming motility on a tryptone swim plate (0.3% agar). (B) Swarming motility on a 0.5% agar plate. (C) Twitching motility on a thin (3-mm) LB plate containing 1% agar. Twitching is observed as a hazy zone of interstitial growth surrounding the surface colony. (D) Staining with crystal violet of cells in twitching zones that remained attached to the polystyrene surface after removing the agar layer and washing with water. (E to H) Light microscopy of the outside of the twitching zone stained with crystal violet. (E and F) S1 and S2 variant cells at a magnification of ×90; (G and H) rafts of S1 and S2 variant cells oriented toward the expending direction of the twitching area at a magnification of ×900.

Flagellar chemotactic response is impaired in the S variants. The fact that all three types of motilities were affected, in addition to the absence of clear chemotactic rings on swim plates, suggested a defect in chemotaxis. The capillary chemotaxis assay showed that the S variants were not significantly attracted to tryptone, a strong chemoattractant. The relative chemotaxis response (ratio of bacterial cell number in the capillary with tryptone to that without tryptone) was between 3.5 and 4 for the L variant and between 1 and 1.5 for the S variants (Fig. 5) A ratio of 2 or more is considered significant (32), confirming that the S variants displayed a defective chemotactic response.

View larger version (39K): [in this window] [in a new window]   FIG. 5.   Comparison of chemotactic responses of L and S variants. Capillary apparatus with or without 0.1% tryptone as a chemoattractant was prepared as described in Materials and Methods and incubated at 37°C for 45 min. The content of the syringe was then plated onto TSA plates for cell enumeration. Error bars represent the standard deviations of duplicates.

Electron microscopy. The high adherence and impaired motility of the S variants hinted at abnormal pili or flagella. Transmission electron microscopy of cells directly sampled from the edge of colonies revealed that the S variants are hyperpiliated (Fig. 6), with the environment of the cells surrounded with pili fragments. Pili were especially abundant in cell aggregates. Bundles formed by the entwinement of numerous pili, some larger than flagella, were frequently observed.

View larger version (86K): [in this window] [in a new window]   FIG. 6.   Transmission electron micrographs of L (A) and S2 (B) variants. Cells were grown overnight on LB agar plates, transferred onto Formvar-coated copper grids, and stained with phosphotungstic acid. Arrows indicate type IV pili. Bars = 0.2 µm.

Expression of various virulence/survival factors is altered in the S variants. We investigated whether there were any differences in virulence/survival factors other than chemotaxis, motility, and piliation. Pyocyanin, the blue phenazine pigment of P. aeruginosa, is an extracellular secondary metabolite with antibiotic activity. As shown in Table 1, the S variants produced three to fivefold more pyocyanin than the L variant. Furthermore, culture supernatants of the S variants contained about 70% more pyoverdine, the primary siderophore of P. aeruginosa, than the L phenotype. The ECR assay unambiguously demonstrated that the S variants excrete less LasB protease (Table 1). Alginate is an exopolysaccharide that is secreted by P. aeruginosa for the establishment of the biofilm matrix. Considering that the S variants formed a biofilm more readily than the L variants, we examined the production of extracellular alginate. In agitated flask cultures at 37°C, there was no difference in the concentration of uronic acids in the supernatant (Table 1). Rhamnolipids are heat-stable hemolysins, displaying surface-active properties, which are coproduced with other extracellular factors (36, 47). The S variants, especially S2, appeared to produce slightly lower concentrations of this biosurfactant than the parent variant (Table 1). Finally, we examined the ability of the variants to survive stress aggression. As shown in Table 1, there was no clear difference in sensitivity to H₂O₂ between the L and S variants in stationary-phase cells. However, when cells from the logarithmic phase of growth were plated, the zone of inhibition caused by H₂O₂ was about 30% smaller for the L variant than for the S variants, indicating hypersensitivity of the latter. Results obtained on TSA plates did not significantly differ from those obtained on medium A agar.

	DISCUSSION

Top Abstract Introduction Materials and Methods Results Discussion References

Colonial phenotypes and type IV pili. Our work describes the isolation and characterization of phenotypic variants (S phenotype) selectively enriched when P. aeruginosa 57RP was cultivated as a biofilm, whereas the parent L phenotype predominates in suspended growth. We also observed that the L variant of P. aeruginosa 57RP produced a surface pellicle enriched in S variants when cultivated in static liquid medium. Under these growth conditions, S variants demonstrated a strong preference for aggregative growth at the air-broth interface. The presence of hydrophobic type 1 fimbriae in E. coli, Salmonella, and Shigella is known to mediate the formation of surface film in nonagitated aerobic cultures (20, 33, 42). Hasman et al. (20) reported that the physical presence of type 1 fimbriae on E. coli K-12 is responsible for the formation of small, convex colonies, whereas the absence of fimbriae is correlated with larger, flat colonies. A similar correlation is commonly used as an indicator of pilus expression in Neisseria gonorrhoeae, which display type IV pili (29, 45). Hyperpiliated mutants of P. aeruginosa have been shown to form small colonies very similar to the S1 variant colonies (2). Thus, both surface film formation and small-colony phenotype are in agreement with hyperfimbriation of our S variants, as also confirmed by transmission electron microscopy (Fig. 6). Observations that the cell surface of the S variants is more hydrophobic and their adhesivity is higher than for the parent L variant are also coherent with the hyperpiliated phenotype.

Motility and chemotaxis. Characterization of P. aeruginosa mutants which lack twitching motility has allowed the identification of about 34 genes involved in type IV pili biogenesis, regulation, and function in twitching (1, 48). Mutation in any of these genes results in nonpiliated cells, with few exceptions such as strains with defects in pilT or pilU, which overexpress surface pili but are incapable of twitching motility (50, 51). The S variants are apparently not directly affected in any of these genes since they actually formed twitching zones at the agar-polystyrene interface, albeit these were smaller than zones formed by the L variants. Another notable exception is pilH, which encodes a homologue of the enteric CheY response regulator. Strains with a defective pilH gene are piliated but form reduced twitching zones, with the presence of donut-shaped swirls at the outer edge of the motile zone (8). We also noticed, especially with S2, many holes and rings of cells reminiscent of the swirls reported by Darzins (8), suggesting that pilH may be affected in the S variants.

Phase variation. Phase variation is a diversity-generating mechanism ensuring that a portion of a bacterial population will be adapted to survive under new environmental conditions (13). Mechanisms regulated by phase variation are essentially stochastic within a population yet at least partially modulated by environmental signals. We observed that under appropriate environmental and growth conditions, cells of each phenotype could switch to the other phenotype at a fairly high frequency, suggesting that the shift between S and L variant phenotypes is regulated by a phase variation mechanism.

S1 is different from S2. Although the S1 form is the more abundant phenotypic small variant that we observed, other morphotypes were obtained when the parental L variant was cultivated on hexadecane, in static cultures, or as a biofilm. The S2 variant displayed many differences with the S1 variant, and most of its phenotypic idiosyncrasies were expressed with more amplitude (Table 2). As demonstrated by the retention of adherence in the twitching zone (Fig. 4D) and in the biofilm kinetic assay (Fig. 2), S2 appears to have an impaired detachment phenotype and to form defective biofilms. The fact that this variant did not build the fine lattice-like network of cells typically found behind the rafts in twitching motility expansion zones (41) may also be related to this defect. These features could be explained by the much lower reversion frequency to the L phenotype displayed by S2. It suggests that the S2 variant is blocked in the S phase; a mutation might impede its ability to undergo phase variation.

Biofilm phenotype. Bacteria in biofilms are phenotypically different from their freely swimming counterpart (5, 38). Our results indicate that the biofilm way of growth selects for a specific phenotypic population that is highly adherent but with reduced motility. Why would chemotactically deficient cells be selected for in biofilms? Chemotaxis is essentially required in environments that are scarce in nutrients. One of the features of biofilms is to provide an environment where nutrients are continuously trapped by the exopolysaccharide matrix and available to the bacteria (5). Obviously, cells inside a biofilm do not require extensive motility until the time they leave to colonize another available surface (49). Mucoid and rough P. aeruginosa strains isolated from cystic fibrosis patients, thus selected for by a biofilm environment (17), lack flagella or are deficient in chemotaxis (16, 30).