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1 School of Medicine, University of Tasmania Clinical School, 43 Collins St, Hobart, Tasmania 7001, Australia
2 School of Biotechnology and Biomolecular Sciences and Centre for Marine Biofouling and Bio-innovation, University of New South Wales, Sydney, NSW 2052, Australia
Correspondence
Sylvia M. Kirov
S.M.Kirov{at}utas.edu.au
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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Present address: School of Biological Sciences, University of Southampton, Southampton SO17 1BJ, UK.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). In the lung, mucus secretions become depleted of paraciliary fluid in the lower respiratory tract, impairing the clearance of inhaled microbes, resulting in chronic infections and an exaggerated inflammatory response that leads to progressive deterioration in lung function and ultimately premature death (Lyczak et al., 2002; Saiman & Siegel, 2004). The opportunistic bacterial pathogen Pseudomonas aeruginosa is the most important agent involved in chronic lung infection in CF patients (Burns et al., 2001; Lyczak et al., 2002). The CF lung environment favours conversion of this organism from a motile, planktonic form to a mucoid, sessile biofilm mode of growth (Costerton et al., 1999; O'May et al., 2006; Worlitzsch et al., 2002). Recognized stages in biofilm development elucidated in experimental model systems include initial attachment, the formation of multicellular structures (‘microcolonies’), and the maturation of microcolonies into thick, three-dimensional structures encased in an exopolymeric matrix and interspersed with water channels (Stoodley et al., 2002). In the CF lung mucus, large aggregates of mucoid P. aeruginosa biofilm communities (growing as per the later stages described in biofilm model systems) have been described (Worlitzsch et al., 2002). In this form the organism becomes impossible to eradicate with existing therapies and is refractory to host defences (Costerton et al., 1999; Hassett et al., 2002).
Although the antibiotic resistance of such infections is well documented, the reasons for it are not fully understood. The mechanisms proposed all reflect the multicellular nature of the biofilm communities. They include the delayed penetration of the agent into the biofilm (exopolymer blocks diffusion or even actively binds agents), the slower growth rate of at least some biofilm bacteria (antibiotics target activities such as cell division and nutrient uptake in growing bacterial cells), and the altered microenvironments (such as anaerobiosis and acidic waste products in the biofilm interior that might directly antagonize the action of an antibiotic or cause physiological changes in the bacteria that lower antibiotic uptakes) (Stewart & Costerton, 2001). Antibiotic-resistant ‘persister’ cell phenotypes that forfeit rapid propagation to ensure population survival in the presence of lethal factors can also be generated in biofilms (Lewis, 2005). More recently, intensive biofilm research of a number of model organisms, including the laboratory strain P. aeruginosa PAO1, in in vitro biofilm models has identified a common biofilm developmental cycle or series of adaptive responses involving an active dispersal mechanism, termed ‘seeding dispersal’, that itself is associated with the generation of genetic diversity in biofilm communities, increasing the chances of survival of the organism (Boles et al., 2004; Hentzer et al., 2005; Koh et al., 2007; Mai-Prochnow et al., 2004; Purevdorj-Gage et al., 2005; Webb et al., 2003).
Seeding dispersal is characterized by ‘hollowing’ in mature microcolonies caused by highly motile cells departing the microcolony interior, leaving behind a stationary cluster of ‘wall’ cells. Hollowing is associated with cell death in a subpopulation of the interior cells. For P. aeruginosa PAO1, these cell death and dispersal events have been linked to the conversion of a prophage to a superinfective lytic form observed at the same time and location as the accumulation of reactive oxygen and nitrogen species (RONS) (Barraud et al., 2006; Webb et al., 2003). Significantly, cells of P. aeruginosa PAO1 released by seeding dispersal exhibit phenotypic and functional diversification. Thus, some biofilm-derived variants show an increased ability to disseminate, whereas others manifest accelerated biofilm formation (Kirisits et al., 2005; Webb et al., 2004). Antibiotic resistance has also been linked to phenotypic variation and biofilms (Drenkard & Ausubel, 2002).
Clearly it is important that more is learnt of the dispersal mechanism and its relevance to clinical isolates if improved methods to deal with chronic P. aeruginosa biofilm infections in CF are to be found. Although seeding dispersal has been well documented in the laboratory strain PAO1 (Purevdorj-Gage et al., 2005), there have been few published studies of the biofilm biology of CF clinical P. aeruginosa strains. Lee et al. (2005) examined a selection of non-mucoid CF isolates and demonstrated that such strains had highly variable biofilm architecture consistent with our and many other published observations that P. aeruginosa isolates with different properties (e.g. decreased motilities and virulence factor expression, increased antibiotic resistance) evolve within the CF respiratory tract (Head & Yu, 2004; Mahenthiralingam et al., 1994; O'May et al., 2006; Oliver et al., 2000). Lee et al. (2005) did not examine mucoid isolates or dispersal in their CF strain biofilms. A mucoid CF strain examined by Purevdorj-Gage et al. (2005) was, however, found to be ‘seeding dispersal negative’, leading to speculation that this mechanism may in fact not be relevant in the clinical setting (Purevdorj-Gage et al., 2005). Here, we expand on our previously published comment on this finding (Kirov et al., 2005) and document experiments that show that seeding dispersal does occur in CF strain biofilms. Moreover, we present evidence that such dispersal events may well be important in the overall persistence of P. aeruginosa in the CF lung by facilitating the release of multiple dispersal cell variants.
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METHODS |
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). The Tasmanian clinical strains of P. aeruginosa (strains 3A, 18A, 32A and 75A) included in this study were isolated from CF sputum samples of chronically infected individuals, as previously described (O'May et al., 2006). The Australian mainland ‘clonal’ CF isolate (strain MC1A) was obtained from the Royal Children's Hospital Melbourne, Australia (Armstrong et al., 2002). These strains were all genotypically distinct by PFGE with restriction enzyme SpeI and random amplified polymorphic DNA (RAPD) assay (Mahenthiralingam et al., 1996). The CF isolates were mucoid on initial isolation and had functional properties (decreased motilities and adherence, impaired biofilm formation in in vitro models especially under anaerobic conditions) typical of Pseudomonas ‘biofilm phenotypes’ isolated from chronically infected individuals (O'May et al., 2006). The strains were further characterized for virulence determinants (LasA protease, LasB elastase and pyocyanin) using methods described elsewhere (Diggle et al., 2002). They were also examined for quorum-sensing (QS) signalling molecules [N-acylhomoserine lactones (AHLs) and 4-hydroxy-2-alkylquinolines (HAQs)]. In brief, these were identified following extraction from overnight LB10 broth cultures, thin-layer chromatography and detection with luxCDABE reporter strains prepared at the Centre for Biomolecular Sciences, University of Nottingham, UK (Diggle et al., 2003; Middleton et al., 2002). Strains were stored at –80 °C in 30 % (v/v) glycerol and subcultured from storage onto Luria–Bertani (LB10) medium [tryptone 10 g l–1, NaCl 10 g l–1, yeast extract 5 g l–1and 15 g l–1 agar], for the experiments below. Bacterial inocula for the biofilm experiments were grown in LB10 broth (
16 h, 37 °C, with shaking) and standardized (OD610) so that an inoculum of 0.5 ml contained 109 bacteria ml–1. Biofilms (continuous-culture flow-cell experiments) were cultivated in M9 minimal medium containing 48 mM Na2HPO4, 22 mM KH2PO4, 9 mM NaCl, 19 mM NH4Cl, 2 mM MgSO4, 100 µM CaCl2 and 5 mM glucose.
Biofilm flow-cell (continuous-culture) system.
Biofilms were grown at room temperature in three-channel flow cells with individual channel dimensions of 0.3x4x40 mm, as described elsewhere (Moller et al., 1998; Webb et al., 2003). Following inoculation with bacteria prepared as described above, the channels were inverted for 2 h with no flow, to allow bacterial adherence. This time was extended beyond the standard 1 h because of decreased adherence abilities of CF strains to glass compared to strain PAO1 (O'May et al., 2006). Media flow (mean velocity of 0.2 mm s–1) was then started and biofilm development followed for up to 8 days, as described below. Three sets of flow cells were run in parallel in each experiment, and duplicate or triplicate channels of each strain were examined in at least two experiments.
Microscopy.
Adhesion and microcolony development of all strains were initially followed daily by bright-field microscopy (Leica DMLB Epifluorescent microscope) and representative fields (minimum five per chamber) were recorded by photography. In brief, individual flow cells were sealed by clamping the in-flow and out-flow tubing to each channel of the flow cell. The flow of M9 medium was then temporarily stopped and individual flow channels were examined directly on the microscope stage. Microcolonies and mature biofilms (days 6–7) were subsequently stained using the BacLight LIVE/DEAD viability kit reagents, SYTO 9 and propidium iodide (Molecular Probes), as described elsewhere (Webb et al., 2003), and examined with a confocal laser scanning microscope (Olympus LSMGB-200). Replicate image stacks (red and green scans) were recorded at 2 µm intervals through the biofilm at random positions in the flow cell. Captured images were subsequently merged to obtain three-dimensional images of live and dead cells within individual microcolonies.
Analysis of colonies from biofilm effluent.
Spent culture medium (1 ml) emerging from individual flow cells was ‘aseptically’ collected into sterile tubes. Serial dilutions (10–2–10–10) of each effluent sample in M9 medium were spread plated onto LB10 agar and detached/dispersed biofilm cells quantified and examined for morphotypic colony variants. Plates were incubated at 37 °C and colonies were observed daily for up to 3 days. Total c.f.u. per ml of each effluent was determined.
Bacteriophage assay.
Effluent samples were also examined for the bacteriophage activity at chosen intervals using a top-layer agar method (Eisenstark, 1967). In brief, aliquots (20 µl) were serially diluted (10–2–10–8) in SM (phage) buffer (180 µl) (Hitch et al., 2004). Phage titres (p.f.u. ml–1) were determined by testing a 10 µl aliquot of each dilution on sectors of LB agar plates overlaid with 3 ml of a top agar of LB10 0.8 % (w/v) agar seeded with an overnight Pseudomonas strain PAO1 LB broth culture. Effluents of selected strains were also tested on agars seeded with CF strain 3A. Phage activity was recorded after overnight incubation of the plates at 37 °C. Each effluent was assayed on duplicate dilutions.
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RESULTS |
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). Thus, after 2 h incubation on the flow-cell glass coverslips, only the Melbourne clonal strain, MC1A, showed comparable adherence to strain PAO1 (data not shown). Strain 32A, an auto-aggregating strain that settled in static broth culture, exhibited small clusters of adherent cells as well as individual attached bacteria. Twenty-four hours post-inoculation, strain PAO1 had formed a layer of cells covering the entire surface. By contrast, the CF strains showed little (strain MC1A) or no change in surface coverage compared to the 2 h time point. After this initial 24 h lag, however, biofilm growth and microcolony development occurred for all strains. By day 5 post-inoculation, each CF strain had its own distinctive and reproducible microcolony architecture. Microcolonies for individual strains varied in their frequency of occurrence, size and structural features as summarized in Table 1 and illustrated in Fig. 1 (bright-field microscopy) and Fig. 2 (confocal microscopy). Some strains (strains 3A, 18A and 32A) formed isolated microcolonies, while for others (strains 75A and MC1A) the microcolonies emerged over time from the mat of cells covering the glass surface, as described in published studies of strain PAO1 (Webb et al., 2003). Microcolony sizes for all strains varied with their stage of development. The ones shown (Fig. 2) and measured (Table 1) were those at the upper size limit and/or (for strain 3A) the ones in which central hollowing (described below) was evident (henceforth referred to as ‘mature microcolonies’). The mean height±SD of such microcolonies for strains 3A, 32A and 75A at days 6–7 (determined by confocal analysis of a combined total of 14 microcolonies of these strains) was 34.5±9.7 µm. Microcolonies of strains 18A (n=6) and MC1A (n=5) were shorter and less variable, at 21.7±0.8 µm and 13.0±1.41 µm, respectively. In the experiments reported here, strain PAO1 grew rapidly, covering the flow cell and forming flat biofilms. Microcolony formation as described in previously published studies was not observed.
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and 2b). By days 5–6, highly active motile bacteria filled the central area (Fig. 3a, arrow). The size of the voids increased with further biofilm development and motile cells were seen breaking out from the outer microcolony structure (Fig. 3b, c, arrows). These observations mirror those of seeding dispersal described for strain PAO1 (Purevdorj-Gage et al., 2005). In contrast, strain 32A had a characteristic encapsulated appearance (large, acellular, amorphous layer surrounding the microcolony) and essentially maintained a dense cellular interior (wall and central cells) over the period of the experiment (Figs 1c, 2e and 3d). Central motile cells were, however, occasionally observed even in the larger microcolonies of this strain. Strain 18A formed numerous, compact microcolonies over the glass surface (Figs 1b and 2c). Generally, these were of smaller size than the microcolonies of strains 3A and 32A. Strain 75A grew as a multilayered, flat biofilm by day 4 from which occasional tall (62.0±13.1 µm) microcolonies emerged from day 5 (Fig. 2a, d). Strain MC1A also initially grew as a flat biofilm that subsequently formed numerous small microcolonies from day 5 onwards (Figs 1d and 2f). Central voids were not apparent by bright-field microscopy for strains 18A, 75A and MC1A at day 5 (e.g. Fig. 1b, d).
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). The spatial organization and degree of bacterial death and lysis varied between strains but was always most marked at the base of the biofilm or microcolony and least apparent at the top of the microcolony. For strain 75A, extensive killing was seen in the centres of its relatively rare microcolonies and appeared identical to that reported for strain PAO1 microcolonies (Fig. 2a, arrow) (Webb et al., 2003). Large patches of cell death (greatest at base) were also observed even in the flat, undifferentiated portions of the biofilm for this strain (Fig. 2d). A similar pattern of lysis was observed in the flat PAO1 biofilms seen in these experiments (not shown). For strains 18A and MC1A, central cell death was also observed within microcolonies. At 7 days this death was more pronounced in strain 18A than in strain MC1A, but central voids were occasionally seen in the larger microcolonies of both strains. Seeding dispersal in strain 18A is evident in Fig. 2(c) (arrows). Strain 3A showed cell death not only from the mature microcolony base upward but also from the microcolony edges inwards as well as central cell death and hollowing (Figs 2b and 3c, arrows). A subpopulation of viable cells was also seen in the central hollow space of the microcolony. Seeding dispersal in association with the cell death and central hollowing was frequently observed for this strain (Fig. 3b, c, arrows). In contrast, the degree of cell death in strain 32A microcolonies was less than that seen in all of the above strains and, although centralized cell death was observed, hollowing and seeding dispersal were rarely observed for this strain in the 8 day experimental period (Fig. 3d).
Phage titre in the effluent of mature CF strain biofilms
Biofilm effluents were examined for viable bacteria (c.f.u.) and bacteriophage (p.f.u.) at days 4, 5, 6 and 7 post-inoculation (one to three experiments). Phage titres were quantified by titration of effluents using a top-layer agar method (usually with strain PAO1 as target). Effluents of strains 3A, 18A and 32A, also examined on CF strain 3A as target, yielded essentially identical phage titres to those obtained on strain PAO1. Between days 4 and 7 phage titres were detected in the effluents from all CF strains as for strain PAO1. For strain PAO1, they exceeded 109 p.f.u. ml–1 throughout (days 4–7). Phage titres of the CF strains were >106 p.f.u. ml–1 at days 4 and 5 post-inoculation and reached levels >109 at day 7, with the exception of strain 32A. For this latter strain, titres were more variable, but they were reproducibly 2–5 logs lower (e.g. <105 p.f.u.) on day 7.
Variant morphotypes in biofilm dispersal effluents
Effluents from strain PAO1 and all CF strain biofilms contained >107 c.f.u. ml–1 (days 5–7 post-inoculation). In contrast to the homogeneous colony morphologies of each of the inoculum strains, at least two colony morphotypes were grown from the biofilm effluents of all strains (Fig. 4). For P. aeruginosa strain PAO1 these were large (>4 mm diameter) colonies, sometimes with an irregular edge, like the inoculum strain, and small-colony (2 mm diameter) variants (SCVs), as previously reported (Webb et al., 2004). The CF strains also showed these two morphotypes in the biofilm effluents. However, the situation was the reverse of that seen with strain PAO1. Despite all strains being mucoid on initial isolation, only strains 3A and 18A retained their mucoidy on subculture, and the other CF biofilm inoculum strains (strains 32A, 75A and MC1A) grew predominantly as small colonies with a translucent edges and yellow centres (Fig. 4b, c, d, e, f, insets). These latter CF inoculum strains gave rise to differing proportions of large-colony variants (LCVs), as well as SCVs. LCVs dominated the effluents of strains 75A and MC1A, but were less common in the effluents from strain 32A (Fig. 4d, e, f). Strains 3A and 18A showed a greater heterogeneity of morphotypic variants, with up to five morphotypes isolated (Table 1; Fig. 4b, c). These strains were also the ones where significant seeding dispersal had been observed, as described above. On subculture on LB10 agar, these biofilm-generated variants were stable and did not revert to the parental morphotypes or convert into other morphotypes over at least two passages (data not shown).
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DISCUSSION |
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, 2004). The flow-cell system allowed for the slower developmental sequence of CF strain biofilms to be followed (8 day experimental period). It was found that the clinical isolates, despite their differences in adhesion and biofilm growth kinetics, and quite distinct microcolony morphologies, exhibited biofilm differentiation events analogous to those described for strain PAO1 (Webb et al., 2003). In particular, it was established that CF strains showed seeding dispersal as the biofilms matured. This active dispersal mechanism, not previously described in clinical strains, also showed a temporal association with the production of lytic bacteriophage and the generation of morphotypic variant dispersal cells as described for strain PAO1. The reproducibility of the differences seen with the CF strains suggests that each of these strains becomes locked into its own pattern of development. Overall, the results confirm that insights gained from studies of strain PAO1 biofilm differentiation and dispersal have clinical relevance. A noteworthy difference, however, was that morphotypic variants recovered from those CF strain biofilms showing marked seeding dispersal were much more heterogeneous than the variants recovered from strain PAO1.
Role of microcolony size, phage induction and cell signalling in seeding dispersal
The observation that seeding dispersal was most commonly seen in CF strains that produced large microcolonies (e.g. strain 3A, with mature microcolony diameters of >100 µm) is consistent with that of Purevdorj-Gage et al. (2005), who reported that a microcolony diameter of 80 µm was required for seeding dispersal in strain PAO1. Microcolonies of the other CF strains studied in this investigation rarely reached this size. In those microcolonies that did reach this size, central motile cells and seeding dispersal were observed. Cell death was, however, seen in the biofilms/microcolonies of all strains as they matured. Although strain PAO1 grew as a flat biofilm in this series of experiments, cell death associated with phage activity and dispersal variant production were nevertheless observed (in patches in the biofilm not in discrete microcolonies). Interestingly, CF strain 75A also formed very flat biofilms similar to strain PAO1, although they were slower to develop. It also rarely formed microcolonies in these experiments. However, the microcolonies that were occasionally seen with this strain had an identical appearance and pattern of central cell death to those previously reported for strain PAO1 (Webb et al., 2003). Biofilm structural maturation has been shown to be dependent on environmental conditions (Klausen et al., 2003; Shrout et al., 2006). Hence, even minor (uncontrolled) differences in conditions could alter the pattern of biofilm development, resulting in flat biofilms especially for the faster-growing, motile strain PAO1, as is not uncommonly observed (Hentzer et al., 2001; Heydorn et al., 2002).
All CF strains showed bacteriophage activity and the phage titres reflected the occurrence of cell death within the biofilms. For instance, strain 32A, with the least death in microcolonies, had significantly lower phage titres than the other strains. To our knowledge, phage production has not previously been documented in relation to CF strain biofilm dispersal, as here. It is, however, now well established that Pf-like filamentous prophages are common among diverse P. aeruginosa clinical isolates. Pf1-like prophage sequences (detected by PCR with phage-specific primers) were present in 8 of 11 CF isolates obtained from different laboratories worldwide (Webb et al., 2003). Additionally, another recent study found that P. aeruginosa isolates from urinary tract catheters and from CF sputa contained Pf1-like phage sequences in their genomic DNA (Mooij et al., 2007). The fact that phage activity from all CF strains was detected on strain PAO1 suggests that these filamentous phages have a broad host specificity in P. aeruginosa. This is supported by the fact that titres of three CF strains tested on CF strain 3A were identical to the titres obtained on strain PAO1. It is also possible that isolates may contain multiple phages, any one of which could cause plaquing of strain PAO1 (Tan et al., 2007).
Purevdorj-Gage et al. (2005) found that las/rhl-mediated QS was required for seeding dispersal to occur in P. aeruginosa PAO1 and our results also support the conclusion that these processes are linked. The loss of QS signal production is a feature of isolates from chronic infections (Hentzer et al., 2005). It has recently been reported that genetic adaptation in the CF lung over time commonly results in mutations in the lasR gene, the elimination of which results in the loss of many acute virulence determinants (Nguyen & Singh, 2006; Smith et al., 2006). Consistent with these reports, several of the clinical strains (strains 32A, 75A and MC1A) studied here were unable to produce one or more of the QS signal molecules and showed loss of acute virulence determinants. QS-negative (lasR) mutants of P. aeruginosa PAO1 are also reportedly significantly more resistant to cell lysis and death than the wild-type (Heurlier et al., 2005). Thus, the loss of AHL signals could possibly affect the degree of cell death and the evacuation of cells through the microcolony wall for these strains. While there did not appear to be a correlation between QS and phage production, strain 32A, which had the lowest phage titres, was also negative for PQS. Interestingly, PQS has been linked to P. aeruginosa autolysis (phage activity) (D'Argenio et al., 2002). Inactivation of lasR results in the accumulation of the PQS precursor, HHQ (Deziel et al., 2004). Thus, a LasR-regulated balance of HAQs may limit P. aeruginosa autolysis (D'Argenio et al., 2007).
Clinical strain dispersal variants show greater diversity than those of strain PAO1
Biofilm growth has been shown to lead to the emergence of colony morphotypic variants in a variety of species (Kirisits et al., 2005; Koh et al., 2007; Mai-Prochnow et al., 2006; Webb et al., 2004). SCVs produced by strain PAO1 have significant functional differences from the wild-type strain (e.g. enhanced attachment, accelerated biofilm development) (Kirisits et al., 2005; Webb et al., 2004). Further studies are needed to investigate the functional significance of CF strain variants and the exact mechanism(s) by which they are generated. In particular, it will be of interest to determine whether there are CF strain variants that are better adapted to colonization and/or survival (e.g. by contributing to antibiotic resistance) under conditions found in the CF airway. The in vitro observations to date are consistent with the high degree of phenotypic variation widely reported in P. aeruginosa isolates from CF sputa despite the fact that most individuals are colonized with one (or relatively few) genotypic clone(s) of the organism (Foweraker et al., 2005; VanDevanter & Van Dalfsen, 2005). It seems likely that in vivo the generation of biofilm dispersal variants represents a key adaptive response that enables protection against antibiotics and host defences, favours recolonization in new niches and ensures continued biofilm growth in pulmonary infection. This study identified variants by colony morphology but morphologically identical clones may show functional variation (Mai-Prochnow et al., 2006). Indeed, we have preliminary evidence from studies of substrate utilization profiles (Biolog Microplates) that this is the case for the dispersal cells of some CF strains.
There are a variety of mechanisms by which variant production may be facilitated in maturing biofilms. Reversible phenotypic and functional variation (e.g. phase variation and persister cells) has been reported in P. aeruginosa (Deziel et al., 2001; Drenkard & Ausubel, 2002; Lewis, 2005) and phase variation observed for some of the CF strains examined in this study (unpublished observations). Thus, LCVs of strain 32A were motile and expressed virulence factors (LasA, LasB and pyocyanin, as well as QS signal molecules) while SCVs of strain 32A and the 32A inoculum strain were non-motile and did not express these virulence factors. Such phenotypic switching could also be observed after serial passage and other culture manipulations of this and other strains from this study (unpublished data).
Biofilm growth also facilitates heritable diversification through genetic exchange and mutagenesis. Two recognized mechanisms that accelerate the latter are the stress-inducible genes, usually part of the SOS regulon, that generate genetic variability in times of stress such as occurs in ageing biofilms and regulator genes (e.g. mutS of the bacterial DNA mismatch repair system), whose functional loss increases the rate of genetic variability and gives rise to ‘hypermutable/mutator’ strains (Barraud et al., 2006; Moyano et al., 2007; Sanders et al., 2006; Smania et al., 2004). Adaptations will be selected for and may be linked to hypermutator mutations if they confer a selective advantage (Hall & Henderson-Begg, 2006; Montanari et al., 2007). The association between multidrug antimicrobial resistance and hypermutability in chronic CF infection has been well documented (Maciá et al., 2004, 2005; Oliver et al., 2000, 2004). Other common genes targeted in chronic infection include the lasR and mexZ genes (Smith et al., 2006). Loss of lasR confers a growth advantage with particular carbon and nitrogen sources as well as increased β-lactamase activity and resistance to ceftazidime, a widely used β-lactam antibiotic in CF (D'Argenio et al., 2007). Loss of mexZ upregulates the multidrug-efflux pump, MexXY-OprM, that is associated with resistance to the aminoglycoside tobramycin, also commonly used in CF treatment (D'Argenio et al., 2007). While selection of mutS mutations with adaptive responses such as those above may be one mechanism by which CF strains give rise to the greater diversity of biofilm dispersal progeny compared to strain PAO1, there are likely other as-yet-unrecognized genes that are also important in causing hypermutability. A recent study found that the level of mutS mutators in multidrug-resistant epidemic strains (strains likely to be particularly well adapted to CF airway survival) was low (13 %, 2/15) (Kenna et al., 2007).
In summary, this paper reports new insights into how biofilm differentiation and dispersal might contribute to the generation of diversity in CF P. aeruginosa isolates and the persistence and intractable nature of airway infections with this organism. These insights have significant implications for ongoing research and the development of improved treatment strategies for chronic Pseudomonas infections in CF. Early and aggressive therapy to prevent and delay colonization, conversion to mucoidy and biofilm growth, and the selection of mutator strains is clearly desirable, as others have concluded (Kenna et al., 2007). This study, however, suggests that successful treatment is only likely to be accomplished if disease management also includes the targeting of the dominant dispersal variants and/or the mechanisms by which they are generated. Further research into these areas is thus clearly warranted.
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ACKNOWLEDGEMENTS |
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Edited by: P. Cornelis
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Received 16 April 2007;
revised 26 June 2007;
accepted 4 July 2007.
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