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1 Northern Arizona University, Flagstaff, AZ 86001, USA
2 The Center for Biofilms, The University of Southern California, Los Angeles, CA, USA
3 The Institute for Genomic Research, Bethesda, MD, USA
Correspondence
Jeff G. Leid
Jeff.Leid{at}nau.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; Spencer, 2003). Recently, B. anthracis received notoriety through the bioterrorism events of 2001. Despite the emphasis on its role as an agent of bioterrorism or biological warfare, anthrax has been, and continues to be, an important global disease of wildlife and livestock. Interestingly, the environmental bacterium Bacillus subtilis, the pathogen Bacillus cereus, and many other Bacillus species closely and distantly related to B. anthracis have been identified in the biofilm mode of life in a variety of settings (Ren et al., 2004; Branda et al., 2004; Stanley et al., 2003; Lopez et al., 2006).
Biofilms are assemblages of microorganisms that are irreversibly associated with a surface and enclosed in a matrix of extracellular materials (Costerton et al., 1999). Biofilms possess several characteristics that are critical to bacterial virulence, pathogenesis and environmental survival. For example, biofilms have significantly higher resistance to antibiotics and the host immune system than their single-cell planktonic counterparts (Donlan, 2000; Donlan & Costerton, 2002; Stewart & Costerton, 2001; Leid et al., 2002, 2005). In the clinical setting, biofilms are responsible for many chronic diseases, such as osteomyelitis, infectious kidney stones, bacterial endocarditis and cystic fibrosis airway infections (Parsek & Singh, 2003). In the natural world, this general antibiotic resistance likely has survival advantages, as a community of organisms can withstand antibiotic onslaught from its neighbours. One of the hallmarks of microorganisms living as a biofilm is the production of an extracellular polymeric substance (EPS) that is often composed of polysaccharides, proteins, enzymes and even DNA (Steinberger & Holden, 2005; Whitchurch et al., 2002; Donlan, 2002; Parsek & Singh, 2003; Costerton et al., 1999). Although the detailed function of the EPS in biofilms is still elusive, likely because each biofilm EPS is unique, it is a universal trait of microorganisms living in the biofilm mode of growth.
One of the best-characterized biofilms within the Bacillus genus is that of B. subtilis (Ren et al., 2004; Branda et al., 2004; Stanley et al., 2003). This common environmental organism has been phenotypically and genetically characterized in the biofilm mode of growth by a number of different investigators (Morikawa, 2006). Additionally, biofilms have been associated with B. cereus, and may represent an important mechanism for B. cereus survival in an agricultural setting (Auger et al., 2006; Rasko et al., 2005; Ehling-Schulz et al., 2004). While these and other Bacillus species have been shown to form biofilms (Morikawa, 2006; Branda et al., 2004, 2006; Ryu & Beuchat, 2005; Shi et al., 2004; Hsueh et al., 2006, Rasko et al., 2005; Donlan, 2002; Auger et al., 2006; Ehling-Schulz et al., 2004; Chu et al., 2006; Lopez et al., 2006), there are no published reports on B. anthracis biofilm formation. Here, we investigated the phenotypic and functional basis of biofilm formation, development and maintenance in B. anthracis over the course of a 168 h culture under static and laminar-shear conditions. At each time point, B. anthracis biofilms were examined for: (1) phenotypic characteristics, such as initial attachment, microcolony formation and sporulation; and (2) functional characteristics associated with antibiotic resistance. We report herein that B. anthracis readily forms biofilms that are more resistant to antibiotics than are planktonic cells. In addition, during biofilm development and maturation, B. anthracis expresses several phenotypes under static and laminar-shear conditions.
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METHODS |
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Growth of B. anthracis.
B. anthracis Sterne was cryogenically stored and aliquots of the stock strain were used for all experiments. The culture was streaked onto BHIA and incubated at 37 °C in the presence of 5 % CO2 for 24 h. After incubation, one isolated colony was inoculated into 10 ml BHIB in a 50 ml plastic conical tube, and incubated at 37 °C with constant agitation overnight. Subsequently, 100 µl overnight culture was inoculated into 10 ml fresh BHIB in a 50 ml plastic conical tube, and incubated at 37 °C with constant agitation for 4 h, until the culture reached the approximate mid-point of the exponential-growth phase, as determined by growth-curve analysis.
Static biofilm formation.
For characterization of the static biofilm phenotype in B. anthracis, the standard Christensen–Kolter–O'Toole microtitre assay was employed (Christensen et al., 1995; O'Toole & Kolter, 1998). The 4 h cultures described above were diluted 1 : 50 in BHIB, and 100 µl culture was transferred into each well of a polystyrene or glass-bottomed (for microscopy) 96-well microtitre plate. The 96-well microtitre plates were covered with Parafilm and incubated at 37 °C in the presence of 5 % CO2 for 8, 24, 48, 72, 144 and 168 h. The medium was replaced every 24 h with fresh BHIB by inverting the plates to remove the old medium and non-adherent cells, and adding 100 µl fresh BHIB to each well under sterile conditions. This procedure allowed for biofilm-specific characterization over time, because the biofilm organisms are attached and the planktonic organisms are discarded (Leid et al., 2005).
Flow cell biofilm formation.
B. anthracis biofilms were grown at 37 °C in three-channel flow cells (Stapper et al., 2004; Bjarnsholt, et al., 2005; Leid et al., 2002) with individual channel dimensions of 1x4x40 mm, supplied at a flow of 3 ml h–1 with 10 % BHIB. The flow system was assembled and prepared as described previously (Stapper et al., 2004; Bjarnsholt et al., 2005; Leid et al., 2002). Cultures for inoculation of the flow channels were prepared as above by inoculating 100 µl stationary-phase B. anthracis into 50 ml plastic conical flasks containing 10 ml BHIB, followed by growth at 37 °C for 4 h under constant rotation. One millilitre of culture at exponential phase was aseptically injected into each flow channel with a 1 ml syringe and a 26
gauge needle. After inoculation, the flow cells were left coverglass-side down at room temperature for 2 h to allow for initial attachment. Subsequently, the flow cell was moved to 37 °C for growth and evaluation of B. anthracis biofilms at 8, 24, 48, 72, 144 and 168 h under laminar shear. Flow was applied and maintained at a constant rate of 3 ml h–1 with a Watson Marlow 205S peristaltic pump. The mean flow velocity in the flow cell was 0.2 mm s–1.
Microscopy.
B. anthracis biofilms statically grown in glass-bottomed 96-well microtitre plates were observed by epifluorescent and confocal microscopy after 8, 24, 48, 72, 144 and 168 h of growth. Phenotypic observations of the biofilms were performed using the green fluorescent nucleic acid dye SYTO 9 (Invitrogen) and a Leica confocal or epifluorescent microscope. In preparation for microscopy, all the planktonic cells were removed by plate inversion, and the biofilm bacteria fixed onto the wells with 100 % ethanol for 15 min. The ethanol-treated biofilms were washed three times with 1 % PBS. After fixation and washing, SYTO 9 was added to the biofilms and the aluminium foil-covered plates were incubated at 4 °C for 30 min. Excess stain was removed by rinsing three times with 1 % PBS. For the flow-cell studies, 1 ml SYTO 9 was applied to each chamber using sterile 1 ml syringes and a 26 gauge needle, and the biofilms were stained for 40 min while covered with aluminium foil at room temperature. After 40 min staining, all chambers were flushed with BHIB at 3 ml h–1 for 10 min. The stained biofilms were observed with a Leica confocal or epifluorescent microscope for 3D structures, at the different time points. Some of the microscopy results demonstrated a haze surrounding the bacterial cells. This was similar to that which is seen with Pseudomonas aeruginosa biofilms, in which extracellular DNA has been identified as part of the EPS (Whitchurch et al., 2002; Steinberger & Holden, 2005). We have also seen this haze with other biofilm organisms, including those that are Gram-positive (unpublished data). However, we are not reporting the definitive identification of EPS within B. anthracis biofilms in this paper because the EPS is still undefined.
Antibiotic resistance studies.
B. anthracis biofilm antibiotic-resistance profiles were obtained using biofilms statically grown for 8, 24, 48, 72, 144 and 168 h in the presence of 5 % CO2 in the microtitre plate assay. Planktonic B. anthracis was grown in BHIB at 37 °C under constant rotation for the same time. Before biofilm challenge, the microtitre plates were quickly inverted, and the non-attached cells (planktonic) in the supernatant fluid discarded into a reservoir containing 50 % bleach. This ensured that only biofilm cells were challenged with antibiotics for these studies. At each time point, the biofilm and planktonic cells were treated with clinically established concentrations of ciprofloxacin (1.5 µg ml–1) and doxycycline (0.22 µg ml–1), with 10 % bleach as a killing control, or with BHIB as a positive control for growth, for an additional 24 h at 37 °C in the presence of 5 % CO2 (biofilms), or under constant rotation (planktonic cells). Planktonic and biofilm cultures were processed for c.f.u. enumeration. The planktonic cultures were centrifuged, the excess media discarded, and the pellets washed with 1 % PBS, before serial dilution and plating on BHIA. The biofilm samples were sonicated in the microtitre plate, and serial dilutions were performed and plated onto BHIA. Plates were incubated at 37 °C in the presence of 5 % CO2 overnight, and the c.f.u. enumerated after 24 h. The c.f.u. of three trials were averaged and plotted on a log scale. Dramatic antibiotic resistance was observed at 144 and 168 h for the biofilm samples; therefore, biofilm and planktonic bacteria were challenged at these time points with increasing concentrations of ciprofloxacin (0.06, 0.6, 6, 12 and 60 µg ml–1) and doxycycline (0.03, 0.3, 3, 6 and 30 µg ml–1). The published MIC of each antibiotic for B. anthracis is underlined. After 24 h incubation in the presence of the antibiotic dilutions, the c.f.u. were determined as described above.
Characterization of sporulation in B. anthracis biofilms.
To evaluate sporulation of B. anthracis during biofilm growth, the biofilms were grown in two different atmospheric conditions, with and without augmentation of 5 % CO2, using the microtitre plate assay. These studies were done because sporulation is part of the natural lifecycle of B. anthracis, and studies have demonstrated that sporulation occurs during biofilm formation in B. subtilis (Lindsay et al., 2005). Sporulation was evaluated at seven different time points: 24, 48, 72, 96, 120, 144 and 168 h. The numbers of vegetative cells and spores were measured to determine the level (percentage) of sporulation during biofilm development by taking 10 µl sonicated biofilm sample, pipetting it onto a Petroff–Hauser counter, and manually counting the spores. Although the spores are heavier than vegetative cells and may settle to the bottom of the counter, this property makes it easier to obtain accurate spore counts, and Petroff–Hauser counting has been utilized for many years to obtain accurate cell counts. The numbers of vegetative cells and spores were enumerated by visual counting under a light microscope at x100 for three different trials. The 120, 144 and 168 h B. anthracis biofilms were diluted 10 times before counting. The number of spores was calculated, compared to the number of vegetative cells, and reported as a percentage of the total number of vegetative cells.
Statistical analysis.
To determine the statistical significance of antibiotic and sporulation data, the planktonic and biofilm samples were compared by the Student's t test at each time point. The data were compared at each time point and P<0.05 was considered to be significant.
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RESULTS |
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). The attached cells exhibited the classical ‘box-car’ phenotype of B. anthracis vegetative cells. However, there were also intertwined ‘networks’ of attached cells at 8 h that served as scaffolding for further biofilm development over time. At 24 h, intricate network structures were seen (Fig. 1a), which expanded at 48 h (Fig. 1a). Initial formation of cell clusters was observed at this time point. Large clusters of cells surrounded by what visually looked like EPS, but could not be confirmed experimentally because the EPS of B. anthracis biofilms has yet to be characterized, were observed at 72 h (Fig. 1a). As biofilm development continued under the static conditions, multiple large cell clusters were observed at 144 h (Fig. 1a) and 168 h (Fig. 1a), with homogeneous coverage of the glass surface observed at 168 h.
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). At 24 h, the B. anthracis biofilm exhibited a uniformly flat structure, representative of surface coverage. This phenotype continued to develop over 48 and 72 h, with varying biofilm peak heights and visible depth to the overall structure. The greatest height of the static biofilms (>30 µm in height) was observed at 144 h, when large clusters of cells were observed. At the end of the growth period, 168 h, the biofilm community was fairly uniform, with densely clustered communities. Crystal violet staining of the static biofilms was performed and confirmed that B. anthracis formed biofilms similar to those of B. cereus and B. subtilis in a PVC microtitre plate assay (data not shown). Nonetheless, our data demonstrate that B. anthracis also binds to glass surfaces, as all the microscopic studies were conducted with glass as a substrate for biofilm formation (Figs 1 and 2).
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) and 48 h (Fig. 2b), microcolony formation was observed, with large colonies (>10 µm) consistently seen at 48 h (Fig. 2b). A clear phenotypic switch was seen at 72 h under shear force, as large macrocolonies (>25 µm) were observed throughout the flow cell, some reaching heights in excess of 70 µm (Fig. 2c). These large macrocolonies were replaced as the biofilms matured with a mix of large cell clusters surrounded by cell monolayers (144 h, Fig. 2d), and eventual establishment of the well-documented mature biofilm structure of macrocolonies surrounded by liquid voids or water channels (168 h, Fig. 2e).
Antibiotic resistance in planktonic and biofilm B. anthracis
To determine if there was a functional consequence of B. anthracis biofilm formation, planktonic and biofilm organisms were challenged with the commonly prescribed antibiotics ciprofloxacin (1.5 µg ml–1) and doxycycline (0.22 µg ml–1) at each time point of biofilm development described above. Planktonic bacteria were highly susceptible to both ciprofloxacin and doxycycline for up to 120 h (data not shown). Interestingly, at 144 and 168 h, the planktonic populations showed resistance to doxycycline and ciprofloxacin, respectively (data not shown). In contrast, B. anthracis biofilm organisms elicited strong antibiotic resistance to the same concentrations of antibiotics, as rapidly as 8 h post-attachment (Table 1). The overall level of resistance to ciprofloxacin and doxycycline remained constant throughout biofilm development and maturation. Strikingly, at 168 h, the B. anthracis biofilm bacteria were resistant to challenge from 10 % bleach (Table 1). Overall, the biofilm organisms were more resistant to doxycycline than ciprofloxacin.
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). However, as reported above, the planktonic cells at 144 h demonstrated resistance to doxycycline across a broad range of concentrations, peaking at 3 µg ml–1, at which initial killing was observed (Fig. 3a). Conversely, even at high concentrations of both antibiotics, ranging from 40 to 150 times greater than common clinical doses, biofilm organisms were resistant to antibiotic challenge (Fig. 3a).
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). The planktonic cells at this time point were, as reported above, more sensitive to ciprofloxacin across the concentration range than they were to doxycycline. However, there was more killing of planktonic than biofilm microorganisms with doxycycline at this time point (168 h, Fig. 3b).
Sporulation of B. anthracis during biofilm development in the presence and absence of 5 % CO2
Previous reports have demonstrated that sporulation occurs within biofilms of B. subtilis (Lindsay et al., 2005, 2006). Additionally, it is clear that sporulation affects antibiotic susceptibility, and that Bacillus species sporulate under conditions of stress, such as nutrient limitation, which commonly occurs in biofilms. Finally, two recent publications have documented biofilm formation and sporulation of Bacillus species in the gastrointestinal tract (Tam et al., 2006; Barbosa et al., 2005). Since one of the hallmarks of this environment is an increased amount of CO2 versus ambient CO2 (approaching 5 %), we tested whether the presence of 5 % CO2 impacted sporulation during B. anthracis biofilm growth. To determine if the presence of CO2 effected sporulation in B. anthracis biofilms, we grew biofilms for 24–168 h, in the presence and absence of 5 % CO2, and microscopically enumerated the number of spores versus vegetative cells. For clarity, we reported the results as the percentage of spores compared to the total number of vegetative cells within the biofilm. In the absence of CO2 (under normal atmospheric conditions), sporulation within the biofilm populations increased with time, and at 168 h, >50 % of the biofilm population was spores (Table 2). In contrast, when biofilms were augmented with 5 % CO2, sporulation in the B. anthracis biofilm was limited to <12 % throughout biofilm development (Table 2).
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DISCUSSION |
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; Branda et al., 2004, 2006; Ryu & Beuchat, 2005; Shi et al., 2004; Hsueh et al., 2006; Rasko et al., 2005; Donlan, 2002; Auger et al., 2006; Ehling-Schulz et al., 2004; Chu et al., 2006; Lopez et al., 2006; Lindsay et al., 2005). Currently, the role of biofilm formation in B. anthracis is poorly characterized, and these data are the first steps in unravelling the functional consequences of biofilm formation in this biothreat and environmentally important pathogen.
The phenotypic characterization of B. anthracis biofilms demonstrated that the causative agent of anthrax is exquisitely capable of forming biofilms under static and shear conditions. Under static conditions, it took at least 48 h for B. anthracis to develop visible biofilm structures (microcolonies), and mature biofilm formation (macrocolonies) was not achieved until >72 h of biofilm growth (Figs 1 and 2). According to the microscopic images of the static biofilms, B. anthracis cells began to irreversibly attach to the glass surface within 8 h of inoculation, and started to form networks consisting of short chains of B. anthracis 24 h after inoculation (Fig. 1). The formation of individual cell clusters in the biofilms continued at 72 h, with mature biofilm development beginning at 120 h (data not shown) and fully visualized at 144 h (Fig. 1). Under static conditions, we did not observe significant changes in the phenotypic characteristics at 168 h, suggesting that B. anthracis forms mature biofilms within 144 h post-attachment (Fig. 1).
When B. anthracis biofilms were grown in the presence of laminar shear in a flow cell, the biofilm phenotype was quite different both in its dynamics and in its overall development. In the flow-cell experiments, we initially did not observe microorganism attachment at 8 h. Under shear conditions, initial cell attachment and biofilm formation were observed at 24 h of static growth, and consisted of small clumps of cells. At 48 and 72 h, the flow-cell phenotype was similar to that observed at 24 h with individual clusters of cells, except that the clusters of cells were larger, with maximal cell-cluster size seen at 72 h. At 144 and 168 h, the flow-cell biofilm phenotypes consisted of many individual macro cell clusters that were surrounded by voids (water channels), and did not contain a monolayer of cells on the glass surface (Fig. 2). This phenotype has commonly been reported for developmentally mature biofilms in flowing conditions, and is often associated with cell-to-cell communication or quorum sensing (Merritt et al., 2003; Stanley & Lazazzera, 2004). We are currently investigating biofilm-specific gene expression of B. anthracis to determine which genetic events are needed for biofilm formation and maturation. These initial studies have been conducted on the Sterne strain, which lacks the pXO2 plasmid. Since this plasmid encodes, among other things, genes related to capsule production, the presence of pXO2 likely impacts the biofilm phenotype. We have conducted a preliminary study with the Ames strain and have confirmed biofilm formation (unpublished results). Collectively, these phenotypic data demonstrate that B. anthracis readily forms biofilms under different shear conditions and exhibits multiple phenotypes during development.
Characteristic of microorganisms growing as biofilms, B. anthracis biofilm cells were resistant to antibiotics (Table 1, Fig. 3). The resistance observed was specific to the biofilm versus planktonic mode of growth at most time points, although late-stationary-phase planktonic cells were moderately resistant to both doxycycline (144 h) and ciprofloxacin (168 h). Strikingly, B. anthracis biofilm resistance was observed as quickly as 8 h post-attachment (Table 1). This suggests, as has been reported in other biofilm studies (Sauer et al., 2002; Landry et al., 2006), that the transition from planktonic to biofilm cells occurs rapidly, and that even early biofilm (<6 h) cells exhibit dramatic antibiotic resistance. Although not extensively tested, these data suggest that B. anthracis biofilm organisms exhibit broad antimicrobial resistance that is not limited to a specific antibiotic family, and demonstrate a functional basis for biofilm formation in B. anthracis.
It has been widely reported that biofilm microorganisms exhibit antibiotic resistance that is often 50–1000 times higher than that in their planktonic counterparts. When challenged under this setting, dramatic antibiotic resistance was still observed. At 144 h, the B. anthracis biofilm organisms were resistant to high doses of both ciprofloxacin and doxycycline (Fig. 3a). This trend was reversed at 168 h, as the highest concentration of both antibiotics killed most of the biofilm cells (Fig. 3b). Presently, the mechanism(s) behind this altered resistance at 168 h is unknown. As stated above, however, preliminary studies are under way in our laboratory to investigate the global gene expression in B. anthracis during biofilm development.
The investigation of sporulation during biofilm development demonstrated that sporulation is regulated during biofilm growth, likely as a result of nutrient limitation and bacterial stress. Sporulation in B. subtilis and B. cereus biofilms is associated with nutrient limitation, a common feature of biofilms (Lindsay et al., 2006). Sporulation during B. anthracis biofilm formation is interesting because both sporulation and biofilm formation are responses to environmental stresses. However, there was a large difference in the number of spores produced in the presence of 5 % CO2. Our data suggest that sporulation and biofilm formation may be linked through a yet-uncharacterized genetic pathway in B. anthracis. Further studies will demonstrate if this is the case in other, well-characterized strains of B. anthracis.
The sporulation data may have a broad impact on potential biothreats and on B. anthracis ecology. For example, B. anthracis biofilms on the surface of water pipeline distribution systems likely form and release spores, which are harder to detect via DNA-based assays than the vegetative form of B. anthracis. However, very little is known about the ecology of the biofilm–spore interaction in B. anthracis. Therefore, more studies of B. anthracis biofilms and sporulation are needed to discover the possible dangers, and to elucidate methods to protect against and prevent spore formation and release in the natural ecology of B. anthracis.
Interestingly, two recent studies investigated spore formers in the gut of poultry and found that many of the microorganisms were within the genus Bacillus, including B. subtilis, B. cereus and a clone that had considerable homology to B. anthracis, and the authors have suggested that a complex lifecycle of spore, vegetative and biofilm cells exists in the gut environment (Tam et al., 2006; Barbosa et al., 2005). The authors further suggested that the biofilm lifestyle may provide a mechanism for bacterial survival in harsh environments. There have been an increasing number of studies that have focused on the biofilm lifestyle in B. subtilis and B. cereus. Most of these studies have focused on deciphering the biofilm-specific genome or proteome, and since these studies have yet to be completed in B. anthracis, a full comparison between biofilm formation and its role in the ecology and subsequent lifecycle of the organism is not yet attainable. Therefore, it is difficult to compare and contrast B. anthracis biofilms with those of other species, except to say that biofilm maturation occurs at approximately the same time (5 days), and that spores are part of the biofilm community. Our studies of biofilm phenotype did not rely solely on crystal violet staining in the context of the 96-well microtitre assay, as many studies have done in the past, therefore, it is hard to place our phenotypic studies in the context of other published studies.
While it is clear that B. anthracis readily forms biofilms, it is not yet clear what role the biofilm mode of growth plays in the ecology and evolution of this pathogen. Of note, it was not the intention of this initial paper to determine the pathogenesis of B. anthracis in the biofilm mode of growth. However, the fact that the microorganisms in these communities are highly resistant to clinically important antibiotics, and that this resistance is not solely related to sporulation, remains an important health question.
Aside from the medical implications, it is extremely likely that the biofilm mode of life plays a role in the overall ecology of B. anthracis in the environment, as do the biofilms of B. subtilis and B. cereus. The study of Ren et al. (2004) on B. subtilis demonstrated that strong correlations exist among biofilm formation, competence and development and sporulation. Although B. anthracis is a monomorphic species, the biofilm lifestyle may play a role in increased gene transfer, resulting in increased genetic diversity and survival under different environmental conditions. Competence and gene transfer are increased in the biofilm mode of growth in other Gram-positive bacteria (Li et al., 2001, 2002). Past genetic studies by our group have demonstrated genetic diversity in B. anthracis as it relates to ecology and distribution (Keim et al., 2000). It is therefore possible that biofilms in the environment may have contributed to some of the genetic diversity that we and others have observed. Continued studies will elucidate more about the genetic mechanisms of biofilm formation, as well as determine the role of biofilms in anthrax ecology.
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ACKNOWLEDGEMENTS |
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Edited by: A. Fouet
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Received 19 October 2006;
revised 19 February 2007;
accepted 21 February 2007.
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