Figures
Abstract
Bacterial exopolysaccharides have always been suggested to play crucial roles in the bacterial initial adhesion and the development of complex architecture in the later stages of bacterial biofilm formation. However, Escherichia coli group II capsular polysaccharide was characterized to exert broad-spectrum biofilm inhibition activity. In this study, we firstly reported that a bacterial exopolysaccharide (A101) not only inhibits biofilm formation of many bacteria but also disrupts established biofilm of some strains. A101 with an average molecular weight of up to 546 KDa, was isolated and purified from the culture supernatant of the marine bacterium Vibrio sp. QY101 by ethanol precipitation, iron-exchange chromatography and gel filtration chromatography. High performance liquid chromatography traces of the hydrolyzed polysaccharides showed that A101 is primarily consisted of galacturonic acid, glucuronic acid, rhamnose and glucosamine. A101 was demonstrated to inhibit biofilm formation by a wide range of Gram-negative and Gram-positive bacteria without antibacterial activity. Furthermore, A101 displayed a significant disruption on the established biofilm produced by Pseudomonas aeruginosa, but not by Staphylococcus aureus. Importantly, A101 increased the aminoglycosides antibiotics' capability of killing P. aeruginosa biofilm. Cell primary attachment to surfaces and intercellular aggregates assays suggested that A101 inhibited cell aggregates of both P. aeruginosa and S. aureus, while the cell-surface interactions inhibition only occurred in S. aureus, and the pre-formed cell aggregates dispersion induced by A101 only occurred in P. aeruginosa. Taken together, these data identify the antibiofilm activity of A101, which may make it potential in the design of new therapeutic strategies for bacterial biofilm-associated infections and limiting biofilm formation on medical indwelling devices. The found of A101 antibiofilm activity may also promote a new recognition about the functions of bacterial exopolysaccharides.
Citation: Jiang P, Li J, Han F, Duan G, Lu X, Gu Y, et al. (2011) Antibiofilm Activity of an Exopolysaccharide from Marine Bacterium Vibrio sp. QY101. PLoS ONE 6(4): e18514. https://doi.org/10.1371/journal.pone.0018514
Editor: Michael Hensel, University of Osnabrueck, Germany
Received: November 18, 2010; Accepted: March 9, 2011; Published: April 7, 2011
Copyright: © 2011 Jiang et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was supported by the National High-tech R&D Program, number 2007AA09Z418 and 2007AA091506, http://www.most.gov.cn/? the National S&T Major Project, number 2009ZX09301-013, http://www.nmp.gov.cn/; the Program for Changjiang Scholars and Innovative Research Team in University, number IRT0944, http://www.dost.moe.edu.cn/; the Special Fund for Marine Scientific Research in the Public Interest, number 201005024?http://www.soa.gov.cn/soa.htm. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Bacterial biofilms are dense aggregates of cell–cell or surface-attached microorganisms encased in a hydrated extracellular polymeric substances matrix of self synthesized [1], [2]. Bacterial pathogens living in biofilm induce persistent chronic infections due to the resistance to antibiotics and host immune system as a result of the diffusion barrier, metabolism slowly, genetic mutant, biofilm specific phenotypes, persistent cells and so on [3], [4]. Compared with the planktonic ones, bacteria within the biofilm state are upward of 1000-times more resistant to conventional antibiotic treatment and host immune responses [5], [6]. Recent estimates suggest that bacterial biofilms are accounting for over 80% of microbial infections in the body [3], [7]. Therefore, searching for compounds or strategies to decrease bacterial biofilm formation or the antibiotics resistance of the pathogens in biofilms, is very essential and useful for the treatment of biofilm-associated disease.
The traditional approach to prevent biofilm formation in vivo is local administration of biocides [8]. However, the bacterial biofilms often persist in the presence of large doses of traditional antimicrobial agents [9]. Recently various approaches were proposed and expected to be effective in directly preventing or eliminating bacterial biofilms. One of common effective strategies is directly targeting at the component of biofilm. For example, the enzymes to degrade extracellular matrix or exopolysaccharide (EPS) were shown not only to inhibit biofilm formation, but also to remove pre-existing biofilms effectively, although the effects depend on the specificity of EPS matrix composition of biofilms [10]–[13].
EPS is a common component of biofilm and its production is an important feature of the mature biofilm [5]. In many bacteria, increased biofilm formation often correlates with increased EPS production. During the process of biofilm formation, by using EPS glycocalyx polymers, bacterial cells initiate the adhesion to the surface and the development of microcolonies [14]–[17]. In addition, EPSs form the matrix that embeds the bacteria, where additional free bacteria can be entrapped [18], [19]. However, few bacterial EPSs were recently found to negatively regulate biofilm formation. Capsular polysaccharide (CPS) transportation protein gene mutant in Vibrio vulnificus [20] or a putative glycosyltransferase gene deletion in Porphyromonas gingivalis [21] was found to decrease the production of EPSs and enhance biofilm formation of themselves. Jaione Valle el. at reported that E. coli group II capsular polysaccharide exerted broad-spectrum biofilm inhibition activity; however, it had no effect on the established biofilms [22]. In addition, P. aeruginosa extracellular products, mainly polysaccharides, were found to disrupt the established S. epidermidis and S. aureus biofilms [23], but biofilms of other pathogens are not involved.
In this study, we showed that an exopolysaccharide A101 purified from culture supernatant of the marine bacterium Vibrio sp. QY101 not only inhibited biofilm formation by a wide range of Gram-negative and Gram-positive bacteria, but also disrupted the established biofilms of some strains. Furthermore, A101-mediated biofilm disruption significantly decreased the minimum biofilm eradication concentration (MBEC) of antibiotics. And the mechanism underlying the antibiofilm effect of A101 was preliminarily investigated. This is the first reported bacterial EPS that exhibits both biofilm formation inhibition activity and pre-existing biofilm disruption activity.
Results
Vibrio sp. QY101 culture supernatant inhibits biofilm formation of P. aeruginosa FRD1
The alginate lyase-producing marine bacterium Vibrio sp. QY101 was isolated from a decaying thallus of Laminaria [24]. Here its culture supernatant was discovered to inhibit P. aeruginosa FRD1 biofilm formation, which was not caused by the original medium (Figure 1A). It has been reported that the acetylated alginate was very important for P. aeruginosa FRD1 biofilm formation [25]. Therefore, we speculated that the biofilm inhibitory activity of the culture supernatant was due to the alginate lyase activity. However, it was found that the purified alginate lyase from Vibiro sp. QY101 had no biofilm inhibitory activity (data not shown), and the culture supernatant still inhibited P. aeruginosa FRD1 biofilm formation after loss of enzyme activity (Figure 1B), demonstrating the active factor was not alginate lyase.
Vibrio sp. QY101 was grown in AM at 25°C for 4 days. Its supernatant was taken to examine the effect on bacterial biofilm formation. P. aeruginosa FRD1 biofilm formed in micro-tubes at 37°C for 24 h. Attached biofilm biomass was determined by measuring crystal violet retained at OD590 nm. Results are the average of six replicates ± SD at least three independent experiments. (A) 1/10 (v/v) supernatant was added in the cultures. (P<0.01). Control: sterile water; Medium: alginate medium; FS: fermentation supernatant. (B) The supernatant was treated to make the alginate lyase inactive before being added in the cultures. Alginate lyase activity was measured by an increase in absorbance at 235 nm of the reaction products.
To elucidate the active component, ethanol precipitation, dialysis and freeze-dried were applied to extract the active factor from the supernatant of Vibrio sp. QY101. Then proteinase K, nuclease and NaIO4, were used to digest the crude extract (crude-A101), respectively. As shown in Figure 2A and 2B, pretreatment crude-A101 with proteinase K and nuclease had no effect on the antibiofilm activity, while pretreatment with NaIO4 resulted in a more than 60% decrease (Figure 2C). It is well known that NaIO4 is able to hydrolyze polysaccharides by oxidizing the carbons bearing vicinal hydroxyl groups and cleaving the C-C bonds [15], [26]. These data suggested that the biologically active component was likely a polysaccharide.
Crude-A101 (150 mg/ml) was respectively treated with proteinase K (1 mg/ml) (A), DNaseI (100 µg/ml) + RNaseA (25 µg/ml) (B) or NaIO4 (20 mM) (C) at 37°C for 12 h, then taken to measure the antibiofilm activity. The strain used in this experiment was P. aeruginosa FRD1. Error bars represent SD of three independent experiments. “Cr-A101”: Crude-A101.
Isolation and purification of the active polysaccharide
The crude polysaccharides were extracted from the supernatant of QY101 and then were further purified by DEAE Sepharose Fast Flow column. The fraction eluted by 0.5 mol/L NaCl contained abundant active sugars (Figure S1A). The active fraction was pooled, dialyzed, lyophilized and was further fractionated on a Sephacryl S-400 HR column eluting with 0.2 mol/L ammonium bicarbonate (Figure S1B). Finally, one active polysaccharide was obtained and named as A101.
IR spectra and chemical compositions of A101
Fourier transform infrared spectra (FT-IR) of A101 revealed typical characteristics of polysaccharides, which displayed a stretching vibration at 3417 cm−1 for the hydroxyl group and a weak C–H stretching band at 2925 cm−1. The characteristic absorption at 1634 cm−1 was also observed, suggesting the presence of C-O bonds, whose symmetrical stretching vibration was at 1416 cm−1. The signals at 1040–1082 cm−1 were attributed to the stretch vibration of C–O and change angle vibration of O–H.
High performance liquid chromatography (HPLC) traces of the polysaccharide hydrolyzates showed monosaccharide components of A101. As shown in Table S1, the predominant monosaccharides of A101 were glucuronic acid (21.47%), galacturonic acid (23.05%), rhamnose (23.90%) and glucosamine (12.15%). Besides the four abundant monosaccharides, mannose, glucose, galactose and fucose were also present in A101 at low level. Additional, high performance gel permeation chromatography (HPGPC) was applied to elucidate the molecular weight (Mw) of the polysaccharide. One independent and symmetrical peak was identified and the average Mw value of A101 is about 546 KDa.
The inhibitory activity of A101 on biofilm formation does not result from reducing bacterial growth
In the microtiter plates biofilm assays, A101 displayed a dose-dependent inhibitory effect on biofilm formation of P. aeruginosa FRD1 and S. aureus RN6390 (Figure 3). In the presence of 100 µg/ml A101, about 75% of biofilm formation of P. aeruginosa FRD1 was inhibited (Figure 3A), and S. aureus RN6390 revealed >90% inhibition (Figure 3B).
Biofilms of P. aeruginosa FRD1 (A) and S. aureus RN6390 (B) were grown in 96-well microplates containing medium at 37°C for 24 h. Medium was mixed with A101 of different concentrations in a does-dependent manner. Results are the average of six replicates ± SD.
To investigate whether the inhibitory effect of A101 on bacterial biofilm formation might due to the direct growth inhibition, growth curve analysis was performed. The results revealed that both P. aeruginosa FRD1 (Figure 4A) and S. aureus RN6390 (Figure 4B) grew a bit faster in the presence of 100 µg/ml A101, indicating that A101 was not bactericidal.
P. aeruginosa FRD1 (A) and S. aureus RN6390 (B) were incubated in fresh medium with or without (Control) 100 µg/ml A101 at 37°C for 24 h with shaking. At regular intervals, 200 µl cultures were taken to measure the absorbance at OD600 nm. Graph show the mean of three experiments; error bars show SD.
The inhibitory activity of A101 on biofilm formation occurs among multiple strains
To determine the spectrum of A101 antibiofilm activity, the effects of A101 on biofilm formation of various strains (P. aeruginosa, E. coli, Actinobacillus actinomycetemcomitans, S. aureus, S. epidermidis, Enterococcus faecalis) were tested using the microtiter plate assay. As indicated in Figure 5, biofilms formation by 12 out of 15 Gram-negative strains (80%) and 7 out of 10 Gram-positive strains (70%) can be inhibited by A101 under static condition, although 4 tested strains' biofilm formation was stimulated (P. aeruginosa ATCC27853 and CS1, S. aureus Col and E. faecalis OG1RF). Furthermore, A101 exhibited prominent prevention effects (≥50%) on 12 out of 25 tested strains (48%). These data demonstrated the antibiofilm activity of A101 occurred across multiple strains of Gram-negative and Gram-positive bacteria.
Biofilms of Gram-negative (A) and Gram-positive (B) bacteria formed in 96-well containing fresh medium with or without (Control) 100 µg/ml A101 at 37°C for 24 h. Levels of crystal violet retained were measured spectrophotometrically (OD590). Results are the average of six replicates ± SD at least three independent experiments. “*”, P<0.05; “**”, P<0.01; “***”, P<0.001.
Although the polystyrene microtiter plate assay is a simple means of testing bacterial biofilm formation, it measures biofilm formation in static cultures but not in a naturally hydrodynamic environment. To better demonstrate the effect of A101 on bacterial biofilm formation, we used a flow cell model that allows the continuous flow of fresh nutrients into a chamber. The GFP-tagged P. aeruginosa FRD1 and S. aureus RN6390 were grown in flow cell in the absence or continued presence of A101, and biofilm was subsequently analyzed using confocal laser scanning microscopy (CLSM). The results showed that A101 treatment dramatically reduced biofilm development of both strains (Figure 6A and 6B). Analysis using COMSTAT software showed that in the presence of 100 µg/ml A101, the total surface-bound biomass of P. aeruginosa FRD1 dropped to less than 5% of that grown in the absence of A101 (Figure 6C Left), while the same treatment resulted in a more than 99% decrease to the surface-bound biomass of S. aureus RN6390 (Figure 6C Right). These findings demonstrated that A101 also had biofilm inhibitory activity in flow cell with dynamic medium that is similar to the models in vivo.
Flow cells irrigated by medium with and without (Control) 100 µg/ml A101 were inoculated with P. aeruginosa FRD1 at 25°C for 2 days (A) or S. aureus RN6390 at 37°C for 24 h (B). Images and structures of biofilms in flow cells were detected by CLSM. CLSM images are representative of three separate experiments. Biomass of biofilm in flow cells were analyzed using COMSTAT software (C). (P<0.001).
A101 disrupts the established bacterial biofilm
While A101 exhibited an inhibitory activity against bacterial biofilm formation, it was of interest to explore whether the established biofilms were also sensitive to A101. This assay was performed by culturing the strains in flow-cell model and irrigated with medium in absence of A101 to allow the establishment of mature bacterial biofilms. Subsequently, the bacterial biofilms were shifted to medium supplemented with A101, and the fate of the bacterial biofilms was followed by CLSM. The results showed that the pre-existing biofilms of P. aeruginosa FRD1 were disrupted severely when treated with 100 µg/ml A101 (Figure 7A Right). Analysis using COMSTAT software suggested that about 85% of mature biofilms were disappeared (Figure 7C Left). Strangely, A101 couldn't eliminate the mature biofilms of S. aureus RN6390 at all (Figure 7B, Figure 7C Right).
Biofilm of P. aeruginosa FRD1 in flow cells was preformed at 25°C for 2 days (A) and S. aureus RN6390 at 37°C for 24 h (B). Then mediums were switched to fresh ones with or without (Control) 100 µg/ml A101 for an additional 12 hours. Images and structures of biofilms in flow cell were detected by CLSM. CLSM images are representative of three separate experiments. Biomass of biofilm in flow cells were analyzed using COMSTAT software (C). (P<0.001).
A101 increases the antibiotics' capability of killing P. aeruginosa FRD1 biofilm
A marker signifying bacterial biofilm production in pathogens is the development of resistance to antibiotics [3], [27]. To investigate whether A101-mediated biofilm disruption increased the antibiotic sensitivity of bacterial biofilm, the effect of A101 on MBEC of antibiotic was examined. P. aeruginosa FRD1 biofilm, similar to previous reports, even at the highest concentration tested (2048 µg/ml) were not killed by amikacin. However, the viability of P. aeruginosa FRD1 biofilm displayed a different antibiotic response in the presence of A101 (100 µg/ml) that addition of amikacin (128 µg/ml) resulted in complete killing of the biofilm (Table 1). Moreover, no synergy was detected between A101 and the used antibiotic on planktonic P. aeruginosa FRD1, because minimal inhibitory concentration (MIC) and minimal bactericidal concentration (MBC) tests of amikacin resulted in the same value in the presence and absence of A101 (data not shown). These results suggested that A101-mediated biofilm dispersion increased the antibiotic's capability of killing bacterial biofilms by indirectly targeting to bacterial cells.
A101 inhibits cell-surface adherence and intercellular adhesion
It is well established that cell-surface interactions are critical for bacterial biofilm formation. To investigate the effects of A101 on the cell surface interactions, the primary adherence to abiotic surfaces assay was performed. The initial attachment of P. aeruginosa FRD1 to surface displayed no difference in the presence (100 µg/ml) and absence of A101 (Figure 8A Up), whereas significant decreases in the adhesion of S. aureus RN6390 were observed when 100 µg/ml A101 was present (Figure 8A Down). This demonstrated that A101 inhibiting celll-surface interactions occurred on S. aureus RN6390, but not on P. aeruginosa FRD1.
(A) Detection the numbers of P. earuginosa FRD1 (Up) or S. aureus RN6390 (Down) attached on the surfaces with (Right) or without (Left) (Control) 100 µg/ml A101. The numbers of adherent cells to the surfaces were detected by phase-contrast microscopy ×600. (B) Observation the formation of aggregates for P. earuginosa FRD1 (Up) or S. aureus RN6390 (Down) in cultures with (Right) or without (Left) (Control) 100 µg/ml A101. Strains were cultivated in medium (M9 (FRD1) or 20% TSB+0.25% glucose (RN6390)) with or without A101 at 37°C for 24 h with shaking, and then observed. The numbers of cells aggregates in cultures were detected by phase-contrast microscopy ×200.
Generally, cell-cell interactions also significantly contribute to bacterial biofilm formation. P. aeruginosa FRD1 and S. aureus RN6390 often grow in an aggregated state in cultures, forming readily observable clumps (the planktonic biofilm) [28] (Figure 8B Left). However, when A101 was present in medium, the aggregates of P. aeruginosa FRD1 and S. aureus RN6390 were powerfully decreased (Figure 8B Right), suggesting that A101 was able to interfere with the cell-cell interactions and inhibit multicellular aggregates formation of the both strains. And to further observe the effects of A101 on the preformed aggregates, A101 was added to the overnight cultures of strains and incubated for 3 hours. The results showed that the percentage of P. aeruginosa FRD1 but not S. aureus RN6390 in the cultures that were present in aggregates were decreased by A101 (data not shown).
Discussion
To our knowledge, the biodiversity of the marine environment and the associated chemical diversity constitute a practically unlimited resource of new bioactive substances [29], and marine diversity is also a potent source of identifying novel antibiofilm compound(s) [30]. In this study, the marine bacterium Vibrio sp. QY101 was found to secret an antibiofilm exopolysaccharide A101.
Disrupting the multicellular structure of bacterial biofilm was proposed one of the most potential strategies for increasing the sensitivity of pathogens in biofilm to antibiotics and host immune systems [31], such as enzymes dissolving the matrix polymers of the biofilm [13], [32], chemical reactions blocking biofilm matrix synthesis, and analogues of microbial signaling molecules interfering with cell-to-cell communication, required for normal biofilm formation [33], [34]. Here, A101 was demonstrated to powerfully disrupt the multicellular aggregates and established biofilm of P. aeruginosa FRD1, and significantly increased the aminoglycosides antibiotics' capability of killing P. aeruginosa FRD1 biofilm. As P. aeruginosa could cause acute and chronic lung infections and result in significant morbidity and mortality in cystic fibrosis [35], the effectiveness of A101 in inhibiting biofilm growth in a majority of P. aeruginosa strains suggest that A101 might be a powerful therapeutic tool in the treatment of P. aeruginosa infections. That biofilm formation of few tested strains was not inhibited by A101 may be related with the complex and probably multifactorial mechanisms involving in biofilms formation [36], [37]. And similar observation has been reported that P. aeruginosa (mainly extracellular polysaccharides) exhibited different effects on biofilm formation of different S. epidermidis strains [23], [38]. In addition, the exopolysaccharide A101 displayed inhibitory activity towards biofilm formation of many Gram-negative and Gram-positive bacteria, including clinic isolates. A101 is potentially superior to other antibiofilm substances in a multispecies biofilm context.
Based on our findings, the antibiofilm activity of A101 is independent of decreasing bacterial viability. Instead, A101 promoted the bacterial growth a little. However, further investigation indicated that strains did not have the ability to utilize the polysaccharide A101 as carbon source, because strains were not able to grow in medium with A101 as the only carbon source (data not shown). And the mechanisms about the A101 promotion bacterial growth are needed to further investigate.
Bacterial extracellular polysaccharides synthesized and secreted by a wide range of bacteria from various environmental habitats are complex and diverse. Some have been proved to involve the pathogenicity [39], [40], while some were found to serve a structural role benefitting the bacterium by providing protection from antimicrobial agents and host defenses [16], [41]–[43], helping nutrient acquisition [44], or promoting adherence to surfaces [45]–[47] and biofilm formation [17], [48]. A101 and other few bacterial EPSs were found to negatively regulate biofilm formation [20]–[23]. As it is known, polysaccharides functions are based on their divergent compositions, biochemical characteristics and structures [20], [49]. Previous studies showed that EPSs in V. cholerae are one important component of its biofilm [50], [51], which primarily contain neutral sugars glucose and galactose [52]. While E. coli II CPS [53] and V. vulnificus CPS [54] with antibiofilm activity were indicative of highly anionic nature. And A101 also displayed high content of uronic acid sugars and electronegative, although its monosaccharides compositions exhibited complexity. These data suggested that the electronegative uronic acid sugars in A101 may play the crucial role for antibiofilm activities.
Cell-surfaces and cell-cell interactions play an important role in bacterial biofilm formation [55]. Many active compounds secreted by bacteria were reported to mediate the interaction (adhesion and deadhesion) between microorganisms and interfaces [49]. And receptor polysaccharides recognized by a complementary protein adhesin were reported to mediate the establishment of productive cell-to-surface and cell-to-cell contacts [56]–[58]. Recent reports demonstrated that polysaccharide-probes have the potential to interfere with adherence and biofilm formation [59]. The E. coli group II CPS was also reported to inhibit biofilm formation not only by weakening cell-surface contacts but also by reducing cell–cell interactions [22]. In this study, A101 was indicated to inhibit or disrupt the interactions of cell-surfaces and cell-cell. It is thus likely that A101-mediated inhibition or disruption cell-surface and cell-cell interactions induced the antibiofilm effects. Moreover, we found that the complete hydrolysis of A101 lost the antibiofilm activity. It is proposed that A101 may contain the motif recognized by adhesins, which caused competitively blocking the recognition of adhesins and receptor polysaccharides on strains surfaces and interfering with the adherence of cell-surface and cell-cell. However, investigation the possible binding of A101 with adhesins would be needed to further understand the mechanism underlying the antibiofilm effects.
A101 was suggested here to inhibit the cells aggregates formation of both S. aureus RN6390 and P. aeruginosa FRD1, but only disrupt the mature biofilm of P. aeruginosa FRD1. And further investigation demonstrated that A101 was only able to disrupt the formed cells aggregates of P. aeruginosa FRD1, but not that of S. aureus RN6390 (data not shown). It has been reported that binding of phage 1P to R. leguminosarum bv. Trifolii involved a reversible binding to EPS and an irreversible binding to the specific lipopolysaccharides receptors [60], [61]. We proposed that the antibiofilm activity of A101 may be involved irreversible binding or adsorption in S. aureus RN6390, and reversible in P. aeruginosa FRD1. Moreover, previous findings that the three-dimensional structure and matrix composition of biofilm were physical barriers to the penetration of antimicrobial agents [3], led us to hypothesis that the different activity against established biofilms of the two strains could also be related with the different architecture and matrix composition of both biofilms: A101 maybe easily penetrate in P. aeruginosa biofilms, but not penetrate in S. aureus biofilms.
To conclude, our findings suggest that A101 is of significance in controlling biofilm-associated infections, even in a multispecies biofilm context, and the antibiofilm activity appears to represent a new function of bacterial EPSs. However, the antibiofilm activity of A101 warrants further investigations of the structure–activity relationships of A101 and the mechanism underlying the antibiofilm effects.
Materials and Methods
Strains and growth conditions
All strains and plasmids used in this study were listed in Table 2. P. aeruginosa was cultured in M9 medium; S. aureus, S. epidermidis, E. faecalis, and A. actinomycetemcomitans were grown in Tryptone Soya Broth (TSB) (Sigma) supplemented with 0.25% glucose; and E. coli was cultured in Luria-Bertani (LB) medium (Sigma). The marine bacterium Vibrio sp. QY101 was cultured in alginate medium (AM) (w/v, 3% NaCl, 0.5% sodium alginate (Sigma), 0.3% KH2PO4, 0.7% K2HPO4, 0.2% (NH4)2SO4, 0.01% MgSO4, 0.01% FeSO4) at 25°C to prepare A101. All the reagents used here are analytical grade.
Plasmids pMRP9-1-GFP (carbenicillin 200 µg/ml) [62] and pSB2019-GFP (chloromycetin 15 µg/ml) [63] were transformed into appropriate strains by electroporation for biofilm formation experiments in flow cell. The software COMSTAT 7.0 was a kind gift from Dr. Arne Heydorn and used to analyze the biofilm parameters [64], [65].
Isolation and purification of A101
Polysaccharides preparation was carried out as previously described [66], [67]. Vibrio sp. QY101 cells were grown in AM with shaking at 25°C for 4 days. The culture supernatant was concentrated under reduced pressure at 40°C, and the EPS was precipitated by adding threefold of the volume of 95% (v/v) ethanol. After keeping at 4°C overnight, the precipitate was dried in a Speed Vac concentrator and dissolved in distilled water followed by dialysis through a 14,000-molecular-weight cutoff membrane. The retained fraction was recovered, concentrated under reduced pressure. After removing proteins by Sevage method, it was lyophilized to obtain the crude polysaccharide. The solution of crude polysaccharide was applied to a DEAE Sepharose Fast Flow column (2.6×50 cm) (Pharmacia) and was eluted by a step gradient of NaCl concentration (0–2 mol/L) at a flow rate of 60 ml/h. Total sugar content of the elution was determined by the phenol-sulfuric acid method. The fractions eluted by 0.5 mol/L NaCl were pooled, desalted and loaded on a Sephacryl S-400 HR column (2.6×100 cm) (Pharmacia) equilibrated with 0.2 mol/L ammonium bicarbonate. The elution was done with the same solution at a flow rate of 12 ml/h, and the active polysaccharide fractions were concentrated under reduced pressure named as A101.
Analysis of the infrared spectra and chemical composition of A101
For IR spectroscopy, as previously described [68], A101 was mixed with KBr powder, ground and then pressed into a 1 mm pellets for FT-IR measurement in the frequency range of 4000–500 cm−1. FT-IR spectra of A101 was measured on a Nicolet Nexus 470 spectrometer.
Total sugar content was determined by the phenol–sulfuric acid method using glucose as the standard [69]. Protein content was measured by the method of BCA. The molecular weight were assessed by HPGPC [68]. The monosaccharide components were determined by HPLC[68]. The standard saccharides used here were glucose, galactose, mannose, rhamnose, xylose, fucose, arabinose, glucuronic acid, galacturonic acid and glucosamine.
Biofilm assay
Static biofilm formation assay was carried out in 96-well microtiter plates (Costar, Cambridge, MA, USA) or mircrotiter tubes as previously described [70], [71]. Briefly, overnight cultures were diluted to about 1×106 cfu/ml with fresh sterile medium, and each well of microtiter plates was filled with 100 µl aliquots of the diluted cultures. The plates were incubated at 37°C for 24 h without shaking for biofilm growth, and non-adherent bacteria were washed by 0.9% (w/v) NaCl for three times. Biofilm was strained by 0.1% (w/v) crystal violet solution for 10 min, washed and then dissolved with 33% (v/v) acetic acid for 10 min. Biofilm mass was determined by measuring the absorbance at 590 nm using a microtiter plate reader (Tecan, Milan, Italy).
Dynamic biofilm formation analysis was carried out using a “once-through” flow cell (4.5 mm×2 mm×35 mm) model [70], [72]. Each cell was injected with 0.8 ml of the diluted culture of the tested strain, and incubated for 40 min without flow. Non-adherent bacteria were then flushed from the flow cell by adjusting the pump to a flow rate (4 ml/h for P. aeruginosa [70], 21 ml/h for S. aureus [72]). CLSM was performed as previously described [73]. The images were constructed with the LSM 5 image browser.
Antibiotics susceptibility assay of bacteria and biofilm
MIC, MBC and MBEC were performed as previously described [74]. MICs of antibiotics were determined using the broth microdilution method. Briefly, the test antibiotic was diluted with MHB medium (Sigma) from 1024 µg/ml to 2 µg/ml by 2-fold serial dilutions. The final concentration of bacterial cultures was about 105 cfu/ml. After incubation at 37°C for 24 h, the absorbance at 600 nm was measured using a microplate reader to assess the cell growth. To assay the MBC, 100 µl cultures was taken from each well in the above MIC experiment, centrifuged and washed three times with PBS and plated on a LB agar plate separately, and bacterial cells were enumerated after incubation at 37°C for 24 h. MBEC™ (MBEC Bio-Products, Edmonton, Canada) assays were performed here to detect tolerance to antibiotics of biofilm bacteria. The device used here is a platform carrying 96 polystyrene pegs that fit in a microtiter plate. For biofilm formation, the device was placed in a tray filled with MHB and cells (∼107 cfu/ml) and was incubated on a tilting shaker, which provides a shearing force, at 37°C for 22 h. After biofilms formed on the pegs, non-adherent bacteria on the pegs were washed from the pegs in a 96-well microtiter plate containing sterile 0.9% (w/v) NaCl. Pegs with the bacterial biofilm were placed into a new microtiter plate with fresh medium and incubated at 37°C for 24 h with antibiotic, which was diluted from 1024 µg/ml to 2 µg/ml. The remaining biofilm was transferred from the pegs into a second 96-well microtiter plate containing fresh broth medium by being sonicated for 5 min in a water bath sonicator. This plate was incubated at 37°C for 24 h. Growth of bacteria in a particular well indicates regrowth of planktonic bacteria from surviving biofilm.
Early bacterial adhesion assay
Early adherence to a polystyrene surface was analyzed as previously described [75] with the following modifications. Overnight cultures of tested strains were grown in medium supplemented with or without A101 at 37°C for 12 h, then adjusted with fresh sterile medium with or without A101 to an optical density at 600 nm of 0.1. Each suspension was added to 24-well microtiter plates (Costar, Cambridge, MA, USA) and incubated at 37°C for 1 h. The 24-well microtiter plates were rinsed gently at least six times with sterile phosphate buffer solution (pH 7.5) and strained by 0.1% crystal violet for 20 min. Adherent bacterial cells were observed by phase contrast microscopy and counted. The results represent the means of four different microscopic fields.
Supporting Information
Figure S1.
Purification of the polysaccharides obtained from marine bacterium Vibrio sp. QY101. (A) A mixture of polysaccharides obtained from the marine bacterium Vibrio sp. QY101 was applied to a DEAE Sepharose Fast Flow column and eluted as described in Method. The fractions containing the active polysaccharides were pooled and named as pre-A101. (B) pre-A101 obtained on DEAE Sepharose Fast Flow was applied to a Sephacryl S-400 HR column and eluted as described in Method. The fractions containing the active polysaccharides were pooled and named as A101.
https://doi.org/10.1371/journal.pone.0018514.s001
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Table S1.
The monosaccharides compositions and the average Mw of A101.
https://doi.org/10.1371/journal.pone.0018514.s002
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Acknowledgments
We thank Keith M. Brady, Dennis E. Ohman, Iñigo Lasa, Thomas K. Wood, Ambrose Cheung, Gary M. Dunny, Di Qu, P J Hill and Pradeep K. Singh for kindly providing strains and plasmids used in this study. We thank Arne Heydorn for the kind gift of COMSTAT software. We thank Guangli Yu and Wenjun Mao for providing direction of techniques and equipments for polysccharides identification.
Author Contributions
Conceived and designed the experiments: WY PJ JL. Performed the experiments: PJ JL FH GD XL. Analyzed the data: WY FH JL PJ GD XL YG. Wrote the paper: WY PJ YG.
References
- 1. Costerton JW, Stewart PS, Greenberg EP (1999) Bacterial biofilms: a common cause of persistent infections. Science 284: 1318–1322.
- 2. Greenberg EP (2003) Bacterial communication: tiny teamwork. Nature 424: 134.
- 3. Davies D (2003) Understanding biofilm resistance to antibacterial agents. Nat Rev Drug Discov 2: 114–122.
- 4. Mah TF, Pitts B, Pellock B, Walker GC, Stewart PS, et al. (2003) A genetic basis for Pseudomonas aeruginosa biofilm antibiotic resistance. Nature 426: 306–310.
- 5. Fux CA, Stoodley P, Hall-Stoodley L, Costerton JW (2003) Bacterial biofilms: a diagnostic and therapeutic challenge. Expert Rev Anti Infect Ther 1: 667–683.
- 6. Hoyle BD, Costerton JW (1991) Bacterial resistance to antibiotics: the role of biofilms. Prog Drug Res 37: 91–105.
- 7. Moreau-Marquis S, Stanton BA, O'Toole GA (2008) Pseudomonas aeruginosa biofilm formation in the cystic fibrosis airway. Pulm Pharmacol Ther 21: 595–599.
- 8. Danese PN (2002) Antibiofilm approaches: prevention of catheter colonization. Chem Biol 9: 873–880.
- 9. Donlan RM, Costerton JW (2002) Biofilms: survival mechanisms of clinically relevant microorganisms. Clin Microbiol Rev 15: 167–193.
- 10. Kaplan JB, Ragunath C, Ramasubbu N, Fine DH (2003) Detachment of Actinobacillus actinomycetemcomitans biofilm cells by an endogenous beta-hexosaminidase activity. J Bacteriol 185: 4693–4698.
- 11. Alkawash MA, Soothill JS, Schiller NL (2006) Alginate lyase enhances antibiotic killing of mucoid Pseudomonas aeruginosa in biofilms. Apmis 114: 131–138.
- 12. Lu TK, Collins JJ (2007) Dispersing biofilms with engineered enzymatic bacteriophage. Proc Natl Acad Sci U S A 104: 11197–11202.
- 13. Iwase T, Uehara Y, Shinji H, Tajima A, Seo H, et al. (2010) Staphylococcus epidermidis Esp inhibits Staphylococcus aureus biofilm formation and nasal colonization. Nature 465: 346–349.
- 14. Wang X, Preston JF, Romeo T (2004) The pgaABCD locus of Escherichia coli promotes the synthesis of a polysaccharide adhesin required for biofilm formation. J Bacteriol 186: 2724–2734.
- 15. Mack D, Fischer W, Krokotsch A, Leopold K, Hartmann R, et al. (1996) The intercellular adhesin involved in biofilm accumulation of Staphylococcus epidermidis is a linear beta-1,6-linked glucosaminoglycan: purification and structural analysis. J Bacteriol 178: 175–183.
- 16. Leid JG, Willson CJ, Shirtliff ME, J. Hassett D, Parsek MR, et al. (2005) The exopolysaccharide alginate protects Pseudomonas aeruginosa biofilm bacteria from IFN-{gamma}-mediated macrophage killing. J Immunol 175: 7512–7518.
- 17. Paul NDanese, Leslie APratt, Kolter R (2000) Exopolysaccharide production is required for development of Escherichia coli K-12 biofilm architecture. J Bacteriol 182: 3593–3596.
- 18. Costerton JW, Lewandowski Z, Caldwell DE, Korber DR, Lappin-Scott HM (1995) Microbial biofilms. Annu Rev Microbiol 49: 711–745.
- 19. Bianciotto V, Andreotti S, Balestrini R, Bonfante P, Perotto S (2001) Extracellular polysaccharides are involved in the attachment of Azospirillum brasilense and Rhizobium leguminosarum to arbuscular mycorrhizal structures. Eur J Histochem 45: 39–49.
- 20. Joseph LA, Wright AC (2004) Expression of Vibrio vulnificus capsular polysaccharide inhibits biofilm formation. J Bacteriol 186: 889–893.
- 21. Davey ME, Duncan MJ (2006) Enhanced biofilm formation and loss of capsule synthesis: deletion of a putative glycosyltransferase in Porphyromonas gingivalis. J Bacteriol 188: 5510–5523.
- 22. Valle J, Da Re S, Henry N, Fontaine T, Balestrino D, et al. (2006) Broad-spectrum biofilm inhibition by a secreted bacterial polysaccharide. Proc Natl Acad Sci U S A 103: 12558–12563.
- 23. Qin Z, Yang L, Qu D, Molin S, Tolker-Nielsen T (2009) Pseudomonas aeruginosa extracellular products inhibit staphylococcal growth, and disrupt established biofilms produced by Staphylococcus epidermidis. Microbiology 155: 2148–2156.
- 24. Song K, Yu WG, Han F, Han WJ, Li JB (2003) Purification and characterization of alginate lyase from marine bacterium Vibrio sp. QY101. Sheng Wu Hua Xue Yu Sheng Wu Wu Li Xue Bao 35: 473–477.
- 25. Hentzer M, Teitzel GM, Balzer GJ, Heydorn A, Molin S, et al. (2001) Alginate overproduction affects Pseudomonas aeruginosa biofilm structure and function. J Bacteriol 183: 5395–5401.
- 26. Mack D, Nedelmann M, Krokotsch A, Schwarzkopf A, Heesemann J, et al. (1994) Characterization of transposon mutants of biofilm-producing Staphylococcus epidermidis impaired in the accumulative phase of biofilm production: genetic identification of a hexosamine-containing polysaccharide intercellular adhesin. Infect Immun 62: 3244–3253.
- 27. Drenkard E, Ausubel FM (2002) Pseudomonas biofilm formation and antibiotic resistance are linked to phenotypic variation. Nature 416: 740–743.
- 28. Schleheck D, Barraud N, Klebensberger J, Webb JS, McDougald D, et al. (2009) Pseudomonas aeruginosa PAO1 preferentially grows as aggregates in liquid batch cultures and disperses upon starvation. PLoS One 4: 1–15.
- 29. Carte BK (1993) Marine natural products as a source of novel pharmacological agents. Curr Opin Biotechnol 4: 275–279.
- 30. You J, Xue X, Cao L, Lu X, Wang J, et al. (2007) Inhibition of Vibrio biofilm formation by a marine actinomycete strain A66. Appl Microbiol Biotechnol 76: 1137–1144.
- 31. Stewart PS, Costerton JW (2001) Antibiotic resistance of bacteria in biofilms. Lancet 358: 135–138.
- 32. Nemoto K, Hirota K, Ono T, Murakami K, Murakami K, et al. (2000) Effect of Varidase (streptokinase) on biofilm formed by Staphylococcus aureus. Chemotherapy 46: 111–115.
- 33. Kolodkin-Gal I, Romero D, Cao S, Clardy J, Kolter R, et al. (2010) D-amino acids trigger biofilm disassembly. Science 328: 627–629.
- 34. Parsek MR, Greenberg EP (2000) Acyl-homoserine lactone quorum sensing in gram-negative bacteria: a signaling mechanism involved in associations with higher organisms. Proc Natl Acad Sci U S A 97: 8789–8793.
- 35. Wagner VE, Iglewski BH (2008) P. aeruginosa Biofilms in CF Infection. Clin Rev Allergy Immunol 35: 124–134.
- 36. Van Houdt R, Michiels CW (2005) Role of bacterial cell surface structures in Escherichia coli biofilm formation. Res Microbiol 156: 626–633.
- 37. Coelho LR, Souza RR, Ferreira FA, Guimaraes MA, Ferreira-Carvalho BT, et al. (2008) agr RNAIII divergently regulates glucose-induced biofilm formation in clinical isolates of Staphylococcus aureus. Microbiology 154: 3480–3490.
- 38. Pihl M, Davies JR, Chavez de Paz LE, Svensater G (2010) Differential effects of Pseudomonas aeruginosa on biofilm formation by different strains of Staphylococcus epidermidis. FEMS Immunol Med Microbiol 59: 439–446.
- 39. Campos MA, Vargas MA, Regueiro V, Llompart CM, Albertí S, et al. (2004) Capsule polysaccharide mediates bacterial resistance to antimicrobial peptides. Infect Immun 72: 7107–7114.
- 40. Pier GB, DesJardin D, Grout M, Garner C, Bennett SE, et al. (1994) Human immune response to Pseudomonas aeruginosa mucoid exopolysaccharide (alginate) vaccine. Infect Immun 62: 3972–3979.
- 41. Begun J, Gaiani JM, Rohde H, Mack D, Calderwood SB, et al. (2007) Staphylococcal biofilm exopolysaccharide protects against Caenorhabditis elegans immune defenses. PLoS Pathog 3: 526–540.
- 42. Sahly H, Keisari Y, Crouch E, Sharon N, Ofek I (2008) Recognition of bacterial surface polysaccharides by lectins of the innate immune system and its contribution to defense against infection: the case of pulmonary pathogens. Infect Immun 76: 1322–1332.
- 43. Vuong C, Jovanka MVoyich, Fischer ER, Braughton KR, Whitney AR, et al. (2004) Polysaccharide intercellular adhesin (PIA) protects Staphylococcus epidermidis against major components of the human innate immune system. Cellular Microbiol 6(3): 269–275.
- 44. Gonzalez JE, York GM, Walker GC (1996) Rhizobium meliloti exopolysaccharides: synthesis and symbiotic function. Gene 179: 141–146.
- 45. Johnson MC, Bozzola JJ, Shechmeister IL, Shklair IL (1977) Biochemical study of the relationship of extracellular glucan to adherence and cariogenicity in Streptococcus mutans and an extracellular polysaccharide mutant. J Bacteriol 129: 351–357.
- 46. Ruas-Madiedo P, Gueimonde M, Margolles A, de los Reyes-Gavilan CG, Salminen S (2006) Exopolysaccharides produced by probiotic strains modify the adhesion of probiotics and enteropathogens to human intestinal mucus. J Food Prot 69: 2011–2015.
- 47. Falcieri E, Vaudaux P, Huggler E, Lew D, Waldvogel F (1987) Role of bacterial exopolymers and host factors on adherence and phagocytosis of Staphylococcus aureus in foreign body infection. J Infect Dis 155: 524–531.
- 48. Rinaudi LV, Gonzalez JE (2009) The low-molecular-weight fraction of exopolysaccharide II from Sinorhizobium meliloti is a crucial determinant of biofilm formation. J Bacteriol 191: 7216–7224.
- 49. Neu TR (1996) Significance of bacterial surface-active compounds in interaction of bacteria with interfaces. Microbiol Rev 60: 151–166.
- 50. Kierek K, Watnick PI (2003) The Vibrio cholerae O139 O-antigen polysaccharide is essential for Ca2+-dependent biofilm development in sea water. Proc Natl Acad Sci U S A 100: 14357–14362.
- 51. Fong JC, Syed KA, Klose KE, Yildiz FH (2010) Role of Vibrio polysaccharide (vps) genes in VPS production, biofilm formation and Vibrio cholerae pathogenesis. Microbiology 156: 2757–2769.
- 52. Yildiz FH, Schoolnik GK (1999) Vibrio cholerae O1 El Tor: identification of a gene cluster required for the rugose colony type, exopolysaccharide production, chlorine resistance, and biofilm formation. Proc Natl Acad Sci U S A 96: 4028–4033.
- 53. Jann K, Jann B, Schmidt MA, Vann WF (1980) Structure of the Escherichia coli K2 capsular antigen, a teichoic acid-like polymer. J Bacteriol 143: 1108–1115.
- 54. Reddy GP, Hayat U, Bush CA, Morris JG Jr (1993) Capsular polysaccharide structure of a clinical isolate of Vibrio vulnificus strain BO62316 determined by heteronuclear NMR spectroscopy and high-performance anion-exchange chromatography. Anal Biochem 214: 106–115.
- 55. O'Toole G KH, Kolter R (2000) Biofilm formation as microbial development. Annu Rev Microbiol 54: 49–79.
- 56. Da Re S, Ghigo JM (2006) A CsgD-independent pathway for cellulose production and biofilm formation in Escherichia coli. J Bacteriol 188: 3073–3087.
- 57. Palmer RJ Jr, Gordon SM, Cisar JO, Kolenbrander PE (2003) Coaggregation-mediated interactions of streptococci and actinomyces detected in initial human dental plaque. J Bacteriol 185: 3400–3409.
- 58. Yoshida Y, Palmer RJ, Yang J, Kolenbrander PE, Cisar JO (2006) Streptococcal receptor polysaccharides: recognition molecules for oral biofilm formation. BMC Oral Health 6(Suppl 1): S12.
- 59. Langille S, Geesey G, Weiner R (2000) Polysaccharide-specific probes inhibit adhesion of Hyphomonas rosenbergii strain VP-6 to hydrophilic surfaces. J Ind Microbiol Biotechnol 25: 81–85.
- 60. By Elzbieta Zajac, Lorkiewicz Z (1983) Role of O-Acetyl groups in the lipopolysaccharide receptor for Rhizobium Phage 1P. J General Microbiol 129: 3171–3176.
- 61. Defives C, Werquin M, Mary P, Hornez JP (1996) Roles of Exopolysaccharides and Lipopolysaccharides in the Adsorption of the Siphovirus Phage NM8 to Rhizobium meliloti M11S Cells. Curr Microbiol 33: 371–376.
- 62. Kaneko Y, Thoendel M, Olakanmi O, Britigan BE, Singh PK (2007) The transition metal gallium disrupts Pseudomonas aeruginosa iron metabolism and has antimicrobial and antibiofilm activity. J Clin Invest 117: 877–888.
- 63. Qazi SN, Rees CE, Mellits KH, Hill PJ (2001) Development of gfp vectors for expression in Listeria monocytogenes and other Low G+C gram positive bacteria. Microb Ecol 41: 301–309.
- 64. Heydorn A, Nielsen AT, Hentzer M, Sternberg C, Givskov M, et al. (2000) Quantification of biofilm structures by the novel computer program COMSTAT. Microbiology 146(Pt 10): 2395–2407.
- 65. Heydorn A, Ersboll BK, Hentzer M, Parsek MR, Givskov M, et al. (2000) Experimental reproducibility in flow-chamber biofilms. Microbiology 146(Pt 10): 2409–2415.
- 66. Hsieh YC, Liang SM, Tsai WL, Chen YH, Liu TY, et al. (2003) Study of capsular polysaccharide from Vibrio parahaemolyticus. Infect Immun 71: 3329–3336.
- 67. Guo S, Mao W, Han Y, Zhang X, Yang C, et al. (2010) Structural characteristics and antioxidant activities of the extracellular polysaccharides produced by marine bacterium Edwardsiella tarda. Bioresour Technol 101: 4729–4732.
- 68. Sun H-H, Mao W-J, Chen Y, Guo S-D, Li H-Y, et al. (2009) Isolation, chemical characteristics and antioxidant properties of the polysaccharides from marine fungus Penicillium sp. F23-2. Carbohydrate Polymers 78: 117–124.
- 69. DuBois M, Gilles KA, Hamilton JK, Rebers PA, Smith F (1956) Colorimetric method for determination of sugars and related substances. Analytical Chemistry 28:
- 70. Musk DJ, Banko DA, Hergenrother PJ (2005) Iron salts perturb biofilm formation and disrupt existing biofilms of Pseudomonas aeruginosa. Chem Biol 12: 789–796.
- 71. Valle J, Toledo-Arana A, Berasain C, Ghigo JM, Amorena B, et al. (2003) SarA and not sigmaB is essential for biofilm development by Staphylococcus aureus. Mol Microbiol 48: 1075–1087.
- 72. Rice KC, Mann EE, Endres JL, Weiss EC, Cassat JE, et al. (2007) The cidA murein hydrolase regulator contributes to DNA release and biofilm development in Staphylococcus aureus. Proc Natl Acad Sci U S A 104: 8113–8118.
- 73. Davies DG, Parsek MR, Pearson JP, Iglewski BH, Costerton JW, et al. (1998) The involvement of cell-to-cell signals in the development of a bacterial biofilm. Science 280: 295–298.
- 74. Brooun A, Liu S, Lewis K (2000) A dose-response study of antibiotic resistance in Pseudomonas aeruginosa biofilms. Antimicrob Agents Chemother 44: 640–646.
- 75. Fournier B, Hooper DC (2000) A new two-component regulatory system involved in adhesion, autolysis, and extracellular proteolytic activity of Staphylococcus aureus. J Bacteriol 182: 3955–3964.
- 76. Douthit SA, Dlakic M, Ohman DE, Franklin MJ (2005) Epimerase active domain of Pseudomonas aeruginosa AlgG, a protein that contains a right-handed beta-helix. J Bacteriol 187: 4573–4583.
- 77. Stapper AP, Narasimhan G, Ohman DE, Barakat J, Hentzer M, et al. (2004) Alginate production affects Pseudomonas aeruginosa biofilm development and architecture, but is not essential for biofilm formation. J Med Microbiol 53: 679–690.
- 78. Gonzalez Barrios AF, Zuo R, Hashimoto Y, Yang L, Bentley WE, et al. (2006) Autoinducer 2 controls biofilm formation in Escherichia coli through a novel motility quorum-sensing regulator (MqsR, B3022). J Bacteriol 188: 305–316.
- 79. Kahl BC, Goulian M, van Wamel W, Herrmann M, Simon SM, et al. (2000) Staphylococcus aureus RN6390 replicates and induces apoptosis in a pulmonary epithelial cell line. Infect Immun 68: 5385–5392.
- 80. Antiporta MH, Dunny GM (2002) ccfA, the genetic determinant for the cCF10 peptide pheromone in Enterococcus faecalis OG1RF. J Bacteriol 184: 1155–1162.