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Journal of Bacteriology, January 2007, p. 119-130, Vol. 189, No. 1
0021-9193/07/$08.00+0     doi:10.1128/JB.00930-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Phenotypic Diversification and Adaptation of Serratia marcescens MG1 Biofilm-Derived Morphotypes{triangledown}

Kai Shyang Koh,1,2 Kin Wai Lam,1,2 Morten Alhede,3 Shu Yeong Queck,1,2,{dagger} Maurizio Labbate,1,2,{ddagger} Staffan Kjelleberg,1,2* and Scott A. Rice1,2

The Centre for Marine Biofouling and Bio-Innovation, The University of New South Wales, Sydney, NSW 2052, Australia,1 The School of Biotechnology and Biomolecular Sciences, The University of New South Wales, Sydney, NSW 2052, Australia,2 Center for Biomedical Microbiology, BioCentrum-DTU, Technical University of Denmark, DK-2800 Kgs. Lyngby, Denmark3

Received 28 June 2006/ Accepted 2 October 2006


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ABSTRACT
 
We report here the characterization of dispersal variants from microcolony-type biofilms of Serratia marcescens MG1. Biofilm formation proceeds through a reproducible process of attachment, aggregation, microcolony development, hollow colony formation, and dispersal. From the time when hollow colonies were observed in flow cell biofilms after 3 to 4 days, at least six different morphological colony variants were consistently isolated from the biofilm effluent. The timing and pattern of variant formation were found to follow a predictable sequence, where some variants, such as a smooth variant with a sticky colony texture (SSV), could be consistently isolated at the time when mature hollow colonies were observed, whereas a variant that produced copious amounts of capsular polysaccharide (SUMV) was always isolated at late stages of biofilm development and coincided with cell death and biofilm dispersal or sloughing. The morphological variants differed extensively from the wild type in attachment, biofilm formation, and cell ultrastructure properties. For example, SSV formed two- to threefold more biofilm biomass than the wild type in batch biofilm assays, despite having a similar growth rate and attachment capacity. Interestingly, the SUMV, and no other variants, was readily isolated from an established SSV biofilm, indicating that the SUMV is a second-generation genetic variant derived from SSV. Planktonic cultures showed significantly lower frequencies of variant formation than the biofilms (5.05 x 10–8 versus 4.83 x 10–6, respectively), suggesting that there is strong, diversifying selection occurring within biofilms and that biofilm dispersal involves phenotypic radiation with divergent phenotypes.


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INTRODUCTION
 
It is now recognized that bacteria in most habitats predominantly live as sessile multicellular consortia, termed biofilms (12), and biofilms form in many environments, including water distribution pipes, river rocks, contact lenses and virtually all indwelling biomedical devices (13). Although biofilm communities are key to many ecologically important processes, such as the biogeochemical cycles, they can also be detrimental, such as in the corrosion of industrial steel pipes, and 60 to 80% of all infections are biofilm based (36). As a result of these biofilm-mediated processes, much emphasis is now given to an improved understanding of the mechanisms of biofilm formation and dispersal.

Recently, phenotypic diversification has emerged as a common theme in biofilms. Significant phenotypic variation has been found to occur within many different biofilm communities (25, 32, 41, 45). Although the biofilm is proposed to provide protection from external stresses, high cell densities inside a biofilm may result in stressful environments characterized by a scarcity of nutrients and oxygen, nonoptimal pH, and accumulation of metabolic by-products (8, 16, 51). Therefore, the ability to evolve and adapt to the different microniches within biofilms is an important survival strategy to thrive in highly variable and adverse environmental conditions (2, 7). In the opportunistic pathogen Pseudomonas aeruginosa, numerous biofilm-derived morphological variants, such as the small-colony variants (SCVs), sticky variants, wrinkly variants, and rugose variants, have been identified (7, 19, 21, 32, 52). Webb et al. (52) demonstrated that the emergence of SCVs was linked to the activity of the Pf4 filamentous phage, which was observed during the dispersal process in P. aeruginosa PAO1 biofilms (53). Furthermore, a dense network of the Pf4 phage was found to be associated with the SCVs (52). Interestingly, these SCV cells exhibited enhanced attachment and biofilm development, and it was proposed that the emergence of the SCVs allows for recolonization of new surfaces during biofilm dispersal. Similarly, Kirisits et al. (32) isolated SCVs of P. aeruginosa, which they termed "sticky" variants, from both laboratory biofilms and cystic fibrosis sputum. It was found that the sticky variants displayed autoaggregative behavior in liquid cultures and exhibited a hyper-biofilm-forming phenotype, potentially facilitating the infection process. The formation of sticky or mucoid variants have been associated with poor outcomes for cystic fibrosis patients (30). In other habitats, phenotypic diversification has been shown to be important for the success of competitive rhizosphere colonization by Pseudomonas fluorescens strains (17, 39).

Serratia marcescens is an emerging pathogen that is responsible for endemic and epidemic nosocomial infections as a result of resistance to many antimicrobial agents (27, 28) and can lead to severe complications, such as Serratia bacteremia (24). This gram-negative bacterium is also implicated in a range of ocular infections such as keratitis, keratoconjunctivitis, conjunctivitis, and endophthalmitis (3, 6, 11, 35). Environmentally, S. marcescens can also cause Cucurbit yellow vine disease in cucurbit crops such as squash (Cucurbita maxima) and pumpkin (Cucurbita pepo) (9), resulting in heavy economic losses. Indeed, S. marcescens MG1 was initially isolated from a rotting cucumber (23). Because biofilms play an important role in diseases of humans and plants, it is important to understand this process in S. marcescens. We have previously characterized the development of the filamentous biofilm of S. marcescens MG1 (34), and it was subsequently shown that under nutrient-limiting conditions S. marcescens MG1 can also form a microcolony-type biofilm (46). In the present study, we have characterized the development of the microcolony-type biofilm of S. marcescens MG1. It was found that the timing and pattern of emergence of six different stable morphological variants emerge in a predictable progression throughout the development of the microcolony biofilm-type of S. marcescens MG1. In addition, phenotypic diversification occurs only in the microcolony biofilm-type and has not to date been identified in the highly porous filamentous biofilm-type, indicating that the internal conditions differ greatly between the two biofilm morphotypes. Comparison of various recolonization traits of the morphological variants, such as motility, attachment, and biofilm formation, showed that these dispersal variants exhibit specialized traits that appear to correlate to the different stages of biofilm development at which they were isolated, suggesting a relationship between biofilm development and functional phenotypic diversification in S. marcescens MG1 biofilms.


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MATERIALS AND METHODS
 
Organisms and culture media. The wild-type S. marcescens strain MG1 (23) was used in the present study. This strain was originally classified as Serratia liquefaciens MG1 but, upon 16S rRNA gene sequencing and phylogenetic analysis, was shown to be S. marcescens and has been reclassified (34a). All S. marcescens strains were routinely grown in M9 minimal medium (47) supplemented with 0.5% (wt vol–1) glucose or on Luria-Bertani (LB) medium (5) supplemented with 1.5% (wt vol–1) agar, unless otherwise indicated. The plate cultures were incubated at 30°C for 3 days, while liquid cultures were incubated with constant agitation at 180 rpm overnight.

Biofilm flow cell experiments. Biofilms were cultivated in glass flow cells, constructed as described previously (46), which were incorporated into the biofilm flow cell system as described by Christensen et al. (10) with some modifications. Overnight cultures of S. marcescens strains were concentrated fourfold with fresh M9 minimal medium supplemented with 0.05% (wt vol–1) glucose and used as inocula for the biofilm experiments. After inoculation of the bacterial concentrate (500 µl) into the glass flow chamber, cells were allowed to incubate for 1 to 1.5 h, after which M9 minimal medium supplemented with 0.05% (wt vol–1) glucose was pumped into the flow cell at a constant rate of 0.15 ml min–1. Three independent experiments were conducted in duplicate for 10 days at room temperature, unless otherwise indicated.

Isolation of phenotypic variants from biofilm effluent. Aliquots of biofilm effluent samples were collected from the 10-day biofilm flow cell experiment on a daily basis. The biofilm effluent samples were vortexed at high speed for 1 min and were serially diluted 10–2 to 10–6 in sterile saline solution. Portions (100 µl) of the serially diluted biofilm effluent samples were plated on LB agar in triplicate and incubated at 30°C for 3 days. Phenotypic dispersal variant colonies were confirmed through visual examination using a Leica ZOOM 2000 dissecting microscope. Frequencies of phenotypic variation of a particular colony phenotype were recorded as the percentage of the total colony counts, i.e., 100 x (the number of variant colonies/the total number of colonies).

Staining and microscopic analysis of biofilms. Biofilms were prepared for confocal scanning laser microscopy by staining with the LIVE/DEAD BacLight viability probe (Molecular Probes, Inc., Eugene, OR), prepared according to the manufacturer's specifications. Two milliliters of working concentration of the LIVE/DEAD BacLight viability probe was pumped into the glass flow cell at a constant rate of 0.15 ml min–1 to minimize the disruption to the biofilms.

Microscopic observation and image acquisition of biofilms was performed by using an Olympus LSMGB20 confocal microscope (Olympus Optical Co., Ltd., Tokyo, Japan) equipped with an argon ion laser at 488 nm for excitation and a 522- to 533-nm band-pass filter or a 605- to 632-nm band-pass for emission. Images were captured by using Olympus LSMGB200 confocal scanning laser microscopy bundled programs and were further processed by using PHOTOSHOP software (version 7.0.1; Adobe Systems, Inc.).

Swarming and swimming assays. Swarming and swimming motility were examined on minimal broth Davis without dextrose (Difco Laboratories, Detroit, MI) supplemented with 0.5% (wt vol–1) Casamino Acids, 0.2% (wt vol–1) glucose, and 0.7% (wt vol–1) or 0.4% (wt vol–1) Bacto agar (Difco Laboratories, Detroit, MI), respectively. Overnight cultures of S. marcescens strains were point inoculated into the center of the agar in 50-mm petri dishes (Sarstedt, Inc.), unless otherwise indicated, and incubated at 30°C for 24 h. Swarming and swimming motility were assessed quantitatively by measuring the diameter of the circular zone formed by the swarming or swimming colony of each strain.

Quantification of bacterial cell attachment and biofilm formation. Biofilm biomass of S. marcescens strains was quantified by a crystal violet staining assay as described previously (42, 43), with some modification. Briefly, seed cultures of S. marcescens strains were prepared by incubation at 30°C for 12 to 16 h, with constant agitation at 50 rpm. For attachment assays, 100 µl of seeding cultures in M9 minimal medium were inoculated into 96-well polystyrene microtiter dishes, while for the biofilm formation assay 1 ml of 1:100 diluted seeding culture in M9 minimal medium supplemented with 0.5% (wt vol–1) glucose was inoculated into 24-well polystyrene microtiter dishes. The microtiter dishes were incubated at 30°C for 2 h without agitation and at 30°C for 24 h with constant agitation at 150 rpm, respectively. The aqueous phase was removed from each well, rinsed three times with phosphate-buffered saline (PBS; pH 7.2), and stained with a 1% (wt vol–1) solution of filtered crystal violet for 20 min at room temperature. The wells were subsequently washed thrice with PBS before the crystal violet-stained biofilms were solubilized in 95% ethanol. The absorbance was measured at 490 nm with a Wallac Victor2 1420 Multilabel counter (Perkin-Elmer, Inc.).

TEM. Bacterial strains used for transmission electron microscopy (TEM) analysis were grown in M9 minimal medium supplemented with 0.5% (wt vol–1) glucose at 30°C overnight, under constant agitation at 50 rpm. Formvar-coated copper 200 mesh grids were floated on drops of the overnight bacterial cell suspension for 1 min, rinsed once with PBS (pH 7.4), and stained with a 2% (wt vol–1) aqueous solution of phosphotungstic acid (pH 7.0) for 20 s. Samples were examined by using a Hitachi H7000 transmission electron microscope at an accelerating voltage of 80 kV. Formvar-coated copper grids were prepared as described by Davison and Colquhoun (15).

Statistical analysis. Data were analyzed by one-way analysis of variance using the SPSS 12.0.2 package for Windows (SPSS, Inc.). Tukey tests provided posthoc comparisons of means.


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RESULTS
 
Characterization of microcolony-type biofilm development in S. marcescens MG1. When grown in M9 minimal medium supplemented with 0.05% (wt vol–1) glucose in a glass flow cell system, S. marcescens MG1 formed a microcolony-type biofilm that followed a consistent pattern of development. Monitoring of the biofilm by confocal microscopy (Fig. 1) indicated that the biofilm developed a limited number of microcolonies by day 2, followed by microcolony maturation and the formation of hollow colonies by days 3 to 4. In contrast to numerous reports in which dead cells could be observed within microcolonies (4, 31, 37, 53), staining using BacLight LIVE/DEAD viability probes (Molecular Probes) clearly revealed the absence of red propidium iodide-stained material in the microcolonies (Fig. 1B). Thus, the microcolonies of S. marcescens appear as empty shells, where the shells are comprised of a thin layer of viable cells. After the formation of large hollow colony structures (>100 µm in diameter), an obvious radiation of biofilm microcolony types was observed (days 5 to 6). This expansion occurred over a very short period, approximately 24 h rather than resulting from a gradual increase in biomass. Subsequently, cell death and biofilm sloughing was observed over the next few days (days 7 to 9), after which biofilm regeneration was noted (day 10). This pattern of biofilm development was consistently observed in at least five independent experiments, each of which were performed in duplicate.


Figure 1
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  		FIG. 1. Biofilm development of S. marcescens MG1 in M9 minimal medium supplemented with 0.05% (wt vol–1) glucose over a 10-day period. (A to F) Confocal microscopic images showing the formation of differentiated microcolonies at day 2 (A); large hollow colony structures at days 3 to 4 (B); biofilm expansion at days 5 to 6 (C); and cell death, biofilm detachment, and biofilm regeneration at days 7 to 10 (D to F). In each panel the top image is the x-y plane, and the arrowhead indicates the position corresponding to the x-z cross-section in the lower image. Magnifications: A, B, and F, x400 (bar, 50 µm); C, x100 (bar, 100 µm); D and E, x200 (bar, 100 µm).

We have previously shown that SwrI, a quorum-sensing (QS) signal synthase that synthesizes N-butanoyl-L-homoserine lactone (20), is required for normal biofilm development into a highly differentiated filamentous biofilm in minimal Davis broth without dextrose (Difco Laboratories) supplemented with 0.05% (wt vol–1) glucose and Casamino Acids (34). To determine whether cell-to-cell signaling (quorum sensing) was also required for the normal development of the microcolony-type biofilm, the signal synthase mutant, S. marcescens MG44, was allowed to form biofilms in M9 minimal medium supplemented with 0.05% (wt vol–1) glucose. When grown in M9 minimal medium (data not shown), the swrI mutant formed microcolonies and proceeded through the normal biofilm developmental pathway. This suggests that quorum sensing, which is essential for filamentous biofilm development, is not required for the differentiation and maturation of the microcolony biofilm-type in S. marcescens MG1.

Occurrence of phenotypic variation in the dispersal cell population of S. marcescens MG1 microcolony-type biofilm. Biofilm effluent was sampled daily from the S. marcescens MG1 biofilm over a period of 10 days and plated onto LB agar plates. Six distinct phenotypic variants were identified (Fig. 2). These morphological colony variants have been termed: sticky-smooth variant (SSV); sticky-rough variant (SRV), sticky-rough-umbonate variant (SRUV), smooth-ultramucoid variant (SUMV), non-sticky-smooth variant (NSV), and non-sticky-small-colony-variant (NSCV). For comparison purposes, after 3 days of incubation at 30°C, the S. marcescens MG1 parental wild-type colonies were ~4 mm in diameter, had a "non-sticky-to-touch" colony texture, and had a rough or grainy appearance (Fig. 2A).


Figure 2
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  		FIG. 2. S. marcescens MG1 phenotypic variants isolated from biofilm effluent. All colonies were grown on LB agar plates for 3 days at 30°C. (A to G) S. marcescens MG1 wild-type (A), SSV (B), SRV (C), SRUV (D), SUMV (E), NSV (F), and NSCV (G) colonies. Bar, 2.5 mm.

The morphological variants were never isolated from the biofilm on days 1 to 2 but first began to appear on days 3 to 4 at the time when the hollow colonies formed (Fig. 1B). Subsequently, the degree of variation in colony morphology increased with the duration of biofilm growth (Fig. 3). Variation in the cell population of the biofilm effluent (for convenience, this will be referred to as the dispersed or dispersal population or the dispersal cells and defined as the cells that are in the biofilm effluent), reached a maximum percentage of total morphotypes between at days 4 to 9, when biofilm expansion was taking place. During the time period of biofilm development, ca. 50% of the dispersing cell population consisted of phenotypic variants. Interestingly, the timing and pattern of variant formation were consistent. For example, SSV always appeared first at low frequencies (3.85% of the population, Fig. 3) at day 3, and SUMV always appeared at the latter stages (days 8 to 10) of biofilm dispersal. The colony morphotypes were stable when passaged daily in liquid culture for 12 days (data not shown), with the exception of NSCV. Some of the NSCV variants were stable, one of which was selected for subsequent phenotypic characterization. This suggests that a genetic change is involved in variant formation for the stable morphotypes as opposed to transient gene expression. Sequencing of the 16S rRNA gene, between positions 27 and 1492 (Escherichia coli 16S rRNA gene sequence numbering), of the phenotypic variants was performed to ensure that the variants were not contaminants. Based on 16S rRNA gene sequence analysis (data not shown), the phenotypic variants exhibited >99.9% identity to the S. marcescens MG1 16S rRNA gene sequence (GenBank accession number AY498856). Also, the variants only appeared during microcolony biofilm formation in S. marcescens and to date have not been observed during filamentous biofilm growth (data not shown).


Figure 3
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  		FIG. 3. Frequency of phenotypic variation (expressed as the percentage of total colony morphotypes) occurring in the effluent of S. marcescens MG1 biofilm grown in M9 minimal medium supplemented with 0.05% (wt vol–1) glucose over a 10-day period. Colonies of dispersal cells bearing typical wild-type colony morphology and the variant phenotypes SSV ({square}), SRV (Figure 3), SRUV (Figure 3), SUMV (Figure 3), NSV (Figure 3), and NSCV (Figure 3) were identified after biofilm effluent were plated on LB agar plates and incubated for 3 days. The wild-type morphotype (not shown) makes up the remaining percentages on each day. The data presented in this graph were obtained from duplicate flow cells and are representative of five independent experiments.

Phenotypic variation results in extensive variation in motility, attachment, and biofilm formation traits. Based on the observations that the timing of the emergence of the phenotypic variants coincided with the development of hollow colonies, biofilm expansion, and biofilm dispersal (days 3 to 10), five of the variants were investigated for changes in traits involved in surface colonization, such as motility, attachment, and biofilm formation. A colony with the typical wild-type colony morphology, termed "BMG1001," isolated at day 10 of biofilm formation was included for comparative purposes. The data obtained showed that the swimming and swarming motility, attachment to different surfaces, and biofilm formation of BMG1001 were similar to the parental wild type (Table 1) . There was no significant difference between the growth rates of the morphological variants and the wild-type (data not shown).


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  		TABLE 1. Comparison of motility, colonization, and biofilm formation traits between parental wild-type and biofilm phenotypic variants

Motility. Assessment of swimming and swarming motility showed that the NSCV exhibited delayed swarming and slightly reduced swimming motility, whereas the rest of the variants (SSV, SRV, SRUV, and SUMV) were only reduced in swimming motility compared to the wild-type (Table 1). Impaired motility for the small colony variants found in P. aeruginosa biofilms has been linked to abnormal pili or flagella (18, 26). TEM of the variants revealed that all of the variants, with the exception of SUMV, were hyperpiliated (Fig. 4), whereas no pili were observed for MG1 wild-type and BMG1001 cells. SUMV cells displayed reduced numbers of pilus structures that were spaced at regular intervals along the bacterial cell and also expressed fewer flagella (one to two flagella per cell) in contrast to abundant peritrichous flagella observed on wild-type cells. The significantly impaired swimming motility (P < 0.001) observed for SUMV could be explained by the reduced number of flagella. In striking contrast to the rod-shaped MG1 wild-type cells (Fig. 4A), NSCV cells were small coccoids that were approximately 0.5 µm in diameter (Fig. 4F). In comparison, the SCVs from P. aeruginosa are microscopically identical to the parent despite forming strikingly different colonies (52). To our knowledge, this is the first report of a biofilm-isolated morphological colony variant that has a cell morphology different from that of wild-type cells.


Figure 4
Figure 4
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  		FIG.4. Transmission electron micrographs of phosphotungstic acid-stained S. marcescens MG1 wild-type (A and i), S. marcescens MG1 biofilm dispersal variants SSV (B and ii), SRUV (C and iii), SRV (D and iv), SUMV (E and v), and NSCV (F and vi) and biofilm dispersal morphotype BMG1001 (G and vii). Panels i to vii represent higher-resolution micrographs of panels A to G, respectively. Black arrows indicate flagellar structures, while the white arrows indicate pili-like structures. Magnifications: A to G, x40,000 (bar, 500 nm); i to vii, x100,000 (bar, 200 nm).

Attachment and biofilm formation. To determine the effects of phenotypic variation on the capacity to colonize and form biofilms on surfaces, the attachment to different surfaces and biofilm formation by the five variants in microtiter plates was measured. Crystal violet staining assays showed that NSCV and SUMV did not attach significantly to either 96-well polystyrene plates (hydrophobic) or tissue culture-treated 96-well polystyrene plates (hydrophilic) and failed to form significant biofilms in 24-well polystyrene plates after 24 h of incubation (Table 1). In contrast, SSV and SRV attached at wild-type levels but formed two- to threefold more biofilm (Table 1), despite showing no differences in growth rates (data not shown).

Phenotypic variation affects differential biofilm formation in flow cells systems. The extensive variation in biofilm formation in a batch system (Table 1) and the unusual biofilm expansion observed for the parental wild-type biofilm within the flow cell (Fig. 1) suggested that the variants were likely to exhibit unique biofilm phenotypes. Using confocal microscopy, we compared and characterized the biofilms formed individually by the five morphological variants in glass flow cell systems, including BMG1001, which displayed the typical wild-type colony morphology. Figure 5 shows the different biofilms formed by BMG1001 and the five variants when cultivated in M9 minimal medium supplemented with 0.05% (wt vol–1) glucose. At day 1, all variants formed either small cell clusters or microcolonies, a finding consistent with microcolony biofilm type development (data not shown). However, extensive differences in biofilm formation by the variants were observed by days 2 to 3. Phenotypic variants SSV, SRV, SRUV, and SUMV formed copious amounts of biofilm which were >100 µm thick, whereas NSCV biofilms were initially composed of differentiated cell chains (Fig. 5E) that coaggregated to form compact irregular microcolony-like structures by days 4 to 5 in M9 minimal medium supplemented with 0.05% (wt vol–1) glucose (data not shown). The SSV biofilms were comprised of large microcolonies (>100 µm in diameter), most of which had hollow interiors (Fig. 5A). The SRV, SRUV, and SUMV biofilms, on the other hand, were generally undifferentiated, forming thick flat biofilms (Fig. 5B to D). Interestingly, the copious amounts of biofilm observed for SRUV and SUMV in flow cell systems contradicts the biofilm formation profile observed in batch biofilm studies, since crystal violet staining of microtiter plates indicated that SRUV and SUMV formed significantly less biofilm than the parental wild-type (Table 1). Similar to the parental MG1 wild-type, BMG1001 formed a limited number of small microcolonies (Fig. 5F) and developed through the typical MG1 microcolony biofilm-type pathway (data not shown).


Figure 5
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  		FIG. 5. Differential biofilm formation of S. marcescens MG1 biofilm dispersal variants in M9 minimal medium supplemented with 0.05% (wt vol–1) glucose. (A to C) Confocal microscopic images of 2-day-old biofilms of SSV (A), SRV (B), and SRUV (C). (D to F) Confocal microscopic images of 3-day-old biofilms of SUMV (D), NSCV (E) and BMG1001 (F). In each panel the top image is the x-y plane, and the arrowhead indicates the position corresponding to the x-z cross-section in the lower image. Magnifications: A to C, x200 (bar, 100 µm); D to F, x400 (bar, 50 µm).

SUMV is a phenotypic variant derived from SSV biofilms. When the SSV biofilm was monitored for 10 days under flow conditions, it was noted that the biofilm structures shifted from large compact microcolonies (day 4 biofilm), which is typical of the SSV, to biofilms comprised of cells that were sparsely distributed within the microcolonies (day 6 biofilm), which is typical of a young SUMV biofilm and an indication of the overexpression of extracellular polymeric substances (Fig. 6A to C). ImageJ (National Institutes of Health) analysis of the SSV biofilms at days 2 and 6 revealed significant differences (P < 0.0001 [Student t test]) in microcolony architecture (90.73 ± 10.00 µm versus 48.54 ± 3.74 µm in diameter, respectively). Analysis of the cells dispersing from the SSV biofilms revealed that the SUMV morphotype could be consistently isolated from the biofilm effluent from day 5 onward (Fig. 6D). Moreover, the timing of appearance of SUMV in the SSV biofilm was consistent with its emergence in the parental wild-type biofilm (5 days after the appearance of SSV), relative to the emergence of SSV (Fig. 3). Furthermore, the frequency of SUMV in the SSV biofilm effluent increased gradually, from 2% (at day 5) to 55% of the dispersing population at day 8. Conversion to the SUMV phenotype appeared to be stable at ca. 50% of the dispersing cell population over the remaining 2 days. No other variants with different colony morphotypes were observed, and the SSV morphotype was never observed to convert into the SUMV when passaged as a planktonic culture or on agar plates.


Figure 6
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  		FIG. 6. Extended flow cell biofilm experiments (10-day-old biofilm) of SSV biofilms revealed a morphological switch to the SUMV biofilm. (A and B) The formation of microcolonies in a 1- to 2-day-old SSV biofilm (A) is followed by the presence of large hollow colonies structures (>100 µm thick) in a 2- to 4-day-old biofilm (B). (C) Sloughing of the biofilm takes place between days 3 to 5, and this was followed by biofilm regeneration at day 5 to 6, resulting in biofilm structural characteristics that resemble a SUMV biofilm. (D) The frequency of phenotypic variation (expressed as the percentage of total colony morphotypes) was determined by plating the biofilm effluent sampled from the SSV biofilm onto LB agar plates daily, and differentiation of SSV (data not shown) and SUMV (Figure 6) morphotypes were identified after 3 days of incubation at 30°C. In panels A to C the top image is the x-y plane, and the arrowhead indicates the position corresponding to the x-z cross-section in the lower image. Magnifications: A and C, x400 (bar, 50 µm); B, x100 (bar, 200 µm).


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DISCUSSION
 
Quorum sensing is not required for microcolony development in Serratia marcescens MG1. This study showed that microcolony biofilm development in S. marcescens MG1 using M9 minimal medium supplemented with 0.05% (wt vol–1) glucose was quorum sensing independent. In contrast, it was previously demonstrated that the swrI mutant grown in minimal broth Davis supplemented with 0.05% (wt vol–1) glucose and 0.05% (wt vol–1) Casamino Acids remained as a flat undifferentiated biofilm and was unable to progress to form highly differentiated cell chain structures observed for the wild type (34). In a recent report, Rice et al. (46) demonstrated that numerous quorum-sensing-dependent traits, such as biofilm formation and swarming motility, can be circumvented by growing the swrI mutant in 0.1X LB medium. Similarly, Davis et al. (14) demonstrated that quorum sensing is important for biofilm development when P. aeruginosa PAO1 was grown in minimal glucose medium, whereas several groups (29, 44) found that there were no marked differences in biofilm structures between the PAO1 wild type and the lasI quorum-sensing mutant. The present study further supports the concept that the role for quorum sensing in biofilm formation is not absolute but is dependent on the culture conditions used and that biofilm structures are similarly medium dependent.

Phenotypic variation of biofilm derived morphotypical variants of S. marcescens MG1. Under the conditions reported here, S. marcescens MG1 biofilm development consistently progressed through a series of stages: formation of microcolonies, emergence of hollow microcolonies, rapid biofilm expansion, cell death, and biofilm detachment. The rapid expansion in biofilm development over a 24-h period in S. marcescens, which has not been described for other organisms, consistently takes place after the appearance of hollow microcolonies (Fig. 1). Analysis of the biofilm effluent resulted in the identification of six colony morphological variants that were stable when passaged on agar plates or in liquid medium. Interestingly, the rapid biofilm expansion at day 4 (Fig. 1C) coincided with the sudden increase in the percentage of morphological variants detected in the biofilm effluent (Fig. 3). The frequencies of variation peaked at ca. 50 to 55% from days 4 to 9, and the biofilm morphotypes observed during biofilm expansion appear to match individual biofilms formed by the morphological variants (Fig. 1C and Fig. 5). Hence, we postulate that rapid biofilm expansion and variant formation are linked. Since we have only selected morphological variants that were easily distinguished from the wild type, the natural frequency of variation in S. marcescens biofilm is likely to be higher. This is supported by the pronounced phenotypic variation observed for Pseudalteromonas tunicata D2 cells derived from colonies of wild-type appearance collected from biofilm effluent (38).

S. marcescens MG1 biofilm variant morphotypes exhibit specialized traits. We investigated several traits that are important for recolonization by dispersal cells: cell surface structures, motility, attachment, and biofilm formation. Using TEM, it was observed that all of the S. marcescens biofilm phenotypic variants, with the exception of the SUMV, displayed hyperpiliated phenotypes (Fig. 4). We found, in contrast to previous studies (18, 26, 33), no clear correlation between hyperpiliation and enhanced attachment or hyper-biofilm-forming ability in the S. marcescens biofilm variants. Either the extensive piliation does not translate into increased attachment or the pili interact with specific receptors and may show preferential adhesion to specific surfaces. For example, piliation in S. marcescens was demonstrated to be important for adherence to biotic surfaces such as epithelial cells but not to abiotic surfaces (34a). Interestingly, all variant morphotypes that had sticky-to-touch colony texture exhibited enhanced biofilm formation in flow cell conditions (Fig. 5), while biofilm-forming ability varied extensively under batch conditions (Table 1). It has been widely reported that the "sticky" variants that were isolated from numerous bacterial species were associated with the hyper-biofilm-forming phenotype (32, 54, 55). It may be that the physiological conditions differ between the batch biofilm and flow cell systems, and we hypothesize that phenotypic variants such as SUMV and SRUV are not well adapted to cope under batch environments in which nutrient and oxygen depletion and the accumulation of undesirable metabolic by-products take place rapidly. It is also possible that biofilm formation as determined by the crystal violet-based detection may be an underestimate due to the excessive extracellular polymeric substances not stained by crystal violet, produced by the variants. In the context of the present study, characterization of these phenotypic variants indicated that they were different from the parental strain in a number of properties, such as swimming, swarming, attachment, and biofilm formation.

Variant formation appears to involve genetic radiation and is increased in frequency within the biofilm. In some organisms, phenotypic variation is an unstable trait, such as the persister cells of P. aeruginosa, where the phenotypic change is likely due to gene regulation. However, the morphological variants described here are stable, suggesting a permanent genetic change. This is supported by the observation that the timing and diversification of phenotypic variants followed a predictable sequence and was strongly correlated with the specific stages of biofilm development. For example, the emergence of the SSV consistently occurred during hollow microcolony formation, and the SUMV consistently emerged when cell death and biofilm detachment occurred (Fig. 3). Furthermore, we demonstrated that SUMV could be derived from SSV biofilms (Fig. 6D), and the timing of the appearance of SUMV is consistent with its appearance relative to SSV in biofilms made by the parental wild-type strain (Fig. 3), suggesting that SUMV is a genetic variant derived from SSV and may share common mutations. Given the specific pattern and timing of variant formation and that the same variants are always observed, we hypothesize that either the genes involved contain hotspots for mutation or the local conditions and stresses within the biofilm change over time, leading to mutations in different genes by separate mechanisms.

Is the formation of variants a strategy for creating dispersal cells or simply a response to stress? The formation of variants from the biofilm has several implications. One possibility, raised above, is that the biofilm represents a stressful environment and the formation of variants is a consequence of an increased mutation rate as a response to the stress. Indeed, a comparison of the frequency of genetic variation between biofilm and planktonic cultures revealed that the diversity and frequency of phenotypic variation were significantly different between the two culture conditions. In planktonic cultures, a low percentage of phenotypic variation was observed (reaching a frequency of 5.05 x 10–8 at day 9 and comprising only two morphological variants) compared to the variation frequencies of 4.83 x 10–6 from biofilms (Fig. 3). Variant formation in planktonic cultures was tested in the same medium as for the biofilms, was followed over a 10-day period, and evaluated both shaken and nonshaken cultures incubated at room temperature. The differences in the frequencies of variation in the variant population suggests that biofilm-specific conditions or a selective pressure that is inherent to biofilms may be required to trigger and maintain phenotypic diversification. This biofilm-specific physiology is supported by a comparison between P. aeruginosa biofilms and planktonic cells, which revealed an overall change in more than 50% of the proteome which was specific to biofilm development (49). Furthermore, the proteomic studies by Sauer et al. (49) showed that there were distinct structural and metabolic changes that could be correlated to different stages of biofilm development, indicating that there are physiological characteristics that are unique to biofilms. Our findings revealed that the generation of phenotypic variants is correlated to microcolony biofilm-type development, since phenotypic variation could not be detected in the highly porous filamentous biofilm-type in S. marcescens. These findings suggest that the internal conditions for the two biofilm types, microcolony and filamentous, may differ dramatically where the microcolony biofilm-type may provide the appropriate conditions (e.g., stress or starvation) required for the induction of variant morphotypes. It should be noted that starvation or stress may not be the only factor required for the formation of variants, since the 10-day-old planktonic cultures would also experience starvation and stress conditions. Perhaps the combination of stress and a biofilm-specific regulon are required for variant formation. One interesting possibility that remains to be resolved, is whether the increased frequency of mutations observed for the biofilm grown cells occur in genes that would alter the cells' ability to colonize different sites or to compete in particular niches. Our data clearly indicate that the variants do differ in several characteristics, such as attachment, swarming motility, and biofilm formation, that are important for surface colonization. In this way, the biofilm life cycle may involve the formation of dispersal cells that differ from the parental strain, and these dispersal cells may be considered as an "insurance policy" (7) for the subsequent occupation of new niches. Clearly, there are several conditions that would need to be met for this to accurately reflect the effects observed. First, the mutations would need to specifically occur in genes that alter surface colonization phenotypes. This is possible if such genes contain hot spots for mutations and hence would be over-represented against the general mutational background. Our data suggest that these variants may occur through such a process because of their high frequency and consistent timing of appearance. Moreover, the formation of the SUMV from the SSV would seem to favor this hypothesis. Second, such mutations in a dispersal population should translate into a selective advantage. This remains to be demonstrated for these specific variants, but it has been demonstrated for other microorganisms that variant formation is important for competitive fitness. For example, the wss gene has been shown to be involved in formation of the "wrinkly spreader" variant of Pseudomonas fluorescens (50), and this variant was less fit when grown in competition on roots (22). Sanchez-Contrares et al. (48) noted for P. fluorescens that the wild-type cells attached to the entire root system, whereas the morphological variants attached in a more specialized fashion and were mostly recovered from either the central part of the root or the last centimeter. Finally, Matz et al. (40) showed that for Vibrio cholerae, the biofilm morphotypes were predominantly rugose, whereas the planktonic cells were smooth, suggesting niche specificity for the two morphotypes. Thus, identification of the mutations involved in variant formation in S. marcescens, their frequency, and fitness under different conditions represent very exciting possibilities for understanding the ecological implications of this process.

Microbial persistence in a stressful environment or in animal hosts has been described as survival in the evolutionary fast track (1, 7). In the present study, we have demonstrated that within a highly controlled environment, microorganisms respond through phenotypic diversification to enhance the adaptive potential of the microbial population. Moreover, specialized traits displayed by different phenotypic variants correlated with specific stages of biofilm development. Studies of such functional diversification and the mechanisms by which it occurs will contribute to a better understanding of microbial evolutionary processes, the adaptability of microbial populations, and how to design biofilm control systems.


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ACKNOWLEDGMENTS
 
We thank Michael Givskov for providing the S. marcescens strains MG1 and MG44 used in this study.

This study was supported by the Australian Research Council and the Centre for Marine Biofouling and Bio-Innovation at The University of New South Wales.


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FOOTNOTES
 
* Corresponding author. Mailing address: The School of Biotechnology and Biomolecular Sciences, The University of New South Wales, Sydney, NSW 2052, Australia. Phone: 61-2-9385-2102. Fax: 61-2-9385-1779. E-mail: S.Kjelleberg{at}unsw.edu.au. Back

{triangledown} Published ahead of print on 27 October 2006. Back

{dagger} Present address: Laboratory of Human Bacterial Pathogenesis, Rocky Mountain Laboratories, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Hamilton, MT 59840. Back

{ddagger} Present address: Department of Chemistry and Biomolecular Sciences, Macquarie University, Sydney, Australia. Back


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Journal of Bacteriology, January 2007, p. 119-130, Vol. 189, No. 1
0021-9193/07/$08.00+0     doi:10.1128/JB.00930-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.



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