• We are sorry, but NCBI web applications do not support your browser and may not function properly. More information
Logo of jbacterPermissionsJournals.ASM.orgJournalJB ArticleJournal InfoAuthorsReviewers
J Bacteriol. Aug 2006; 188(16): 5945–5957.
PMCID: PMC1540058

Membrane Vesicles: an Overlooked Component of the Matrices of Biofilms

Abstract

The matrix helps define the architecture and infrastructure of biofilms and also contributes to their resilient nature. Although many studies continue to define the properties of both gram-positive and gram-negative bacterial biofilms, there is still much to learn, especially about how structural characteristics help bridge the gap between the chemistry and physical aspects of the matrix. Here, we show that membrane vesicles (MVs), structures derived from the outer membrane of gram-negative bacteria, are a common particulate feature of the matrix of Pseudomonas aeruginosa biofilms. Biofilms grown using different model systems and growth conditions were shown to contain MVs when thin sectioned for transmission electron microscopy, and mechanically disrupted biofilms revealed MVs in association with intercellular material. MVs were also isolated from biofilms by employing techniques for matrix isolation and a modified MV isolation protocol. Together these observations verified the presence and frequency of MVs and indicated that MVs were a definite component of the matrix. Characterization of planktonic and biofilm-derived MVs revealed quantitative and qualitative differences between the two and indicated functional roles, such as proteolytic activity and binding of antibiotics. The ubiquity of MVs was supported by observations of biofilms from a variety of natural environments outside the laboratory and established MVs as common biofilm constituents. MVs appear to be important and relatively unacknowledged particulate components of the matrix of gram-negative or mixed bacterial biofilms.

Biofilms are now commonly perceived as the predominant form in which microorganisms exist in natural and human-made environments. In themselves, the increased persistence and recalcitrance demonstrated by biofilm populations have been a cause for concern in many industrial, health care, and domestic household settings. The exact manner by which biofilm populations become more versatile and resilient than their corresponding planktonic counterparts is currently a matter of great interest but is still not entirely resolved. Yet, in this respect the matrix is an indispensable feature of biofilms. It contributes to the hallmark recalcitrant nature of biofilms by preventing desiccation and phagocytosis, allowing for the accumulation of nutrients, and permitting enhanced resistance to certain inimical agents, as well as influencing fundamental parameters such as rheology and mechanical stability (64, 65). It also helps define the architecture and spatial arrangement of cells (29, 48), and the creation of demarcated niches within the biofilm, i.e., microenvironments. The latter facilitate both phenotypic heterogeneity and species segregation through the creation of environmental continua, e.g., oxygen (16, 67) and pH gradients (12, 59). This adds greater opportunity for diversity and adaptability within a single biofilm. In this manner, the matrix contributes to the intrinsic properties of biofilms and accounts for the great research effort into its underlying chemical and physicochemical properties.

Initial studies of the complexity of the biofilm matrix heavily emphasized exopolysaccharides as the major chemical constituent (14, 15), yet there is now a growing appreciation of other molecules such as proteins, lipids, and nucleic acids (11, 64). These may in turn interact to form higher-order structures, e.g., lattices of intertwined molecules, or particulate structures such as pili (fimbriae), flagella, phage, and membrane vesicles (MVs), all of which have been reported within biofilms (8, 10, 18, 21, 27, 38, 57, 62). Yet, in spite of the tremendous effort to understand the contribution of matrix chemistry to matrix properties, our current knowledge of the ultrastructure of the matrix is still limited (32).

In our current work, we turned our efforts towards the investigation of MVs as promising candidate particulate structures of the matrix of gram-negative and mixed bacterial biofilms. MVs are complex and chemically heterogeneous bilayered structures derived from the outer membrane of a wide variety of gram-negative bacteria, crossing a number of genera (reviewed by references 9 and 43). As MVs bleb from the outer membrane, periplasm fills their lumen and is retained there. The contents of the lumen vary according to the physiology of the parent organism and may consist of a variety of periplasmic constituents such as proteases, alkaline phosphatase, lipases, proelastase, autolysins, and toxins (2, 20, 25, 31, 34, 36, 39). Apart from the lumen contents, another important feature of MVs is their surface chemistry, which is similar to that of the outer membrane of the parent cell although differences in the relative stoichiometry do exist (30, 34). As MVs form, they also retain the intrinsic lipid asymmetry of the outer membrane with most of the lipopolysaccharide (LPS) positioned within the outer leaflet of the membrane (9). LPS has been reported as a component of the matrix (22, 23, 66), possessing possible structural roles (66), and apart from the cells, MVs could represent a major source of LPS within the biofilm matrix. Given the amphipathic nature of LPS, it would likely be arranged in either vesicles or micelles. Since the excision of LPS from the outer membrane with the exclusion of other membrane components (such as proteins or phospholipids) would not readily occur, MVs could be preferred LPS structures in biofilms, especially as these are naturally shed from gram-negative bacteria (9).

In the first part of our work, we used transmission electron microscopy (TEM) of embedded and thin-sectioned biofilms to view the internal structure of the biofilm. Once the presence of MVs was confirmed in thin sections, isolated MVs provided an additional means of proof. In the second part of our investigation we assessed biofilms grown using a number of established model systems and different growth conditions. All of the methods confirmed that MVs were present in all of the biofilms that we studied. Work with a variety of non-laboratory-grown biofilms, taken from natural and human-made environments, also revealed the presence of MVs and established MVs as a common biofilm phenomenon. In the third and final part of this study, we isolated and characterized MVs from planktonic and biofilm populations and found quantitative and qualitative differences between the two as well as indications of roles that MVs may play within biofilms. Collectively, the results suggest that MVs are ubiquitous and important particulate constituents of the biofilm matrix of gram-negative and mixed bacterial biofilms.

MATERIALS AND METHODS

Bacterial strain and culture maintenance.

Pseudomonas aeruginosa PAO1 was utilized in the study. Cultures were maintained at −20°C in Trypticase soy broth (TSB; BBL, Becton Dickinson and Company) supplemented with glycerol (10%, vol/vol; Fisher) and on Pseudomonas Isolation Agar (Difco; Becton Dickinson and Company) plates and slopes at room temperature. Escherichia coli K-12, Shewanella oneidensis MR-1, and Azotobacter sp. were also used in our study and were maintained on Trypticase soy agar (TSA) plates (BBL, Becton Dickinson and Company) and as frozen stocks.

Biofilm growth.

Biofilms were grown using different model growth systems: agar plate (TSA; BBL, Becton Dickinson and Company; 90-mm diameter, Fisher Scientific), flow cell (flat-plate flow cell, 1 mm by 10 mm by 40 mm; Biosurfaces, Inc.), silicone tubing (internal diameter, 1.6 mm) (49), drip reactor (substrata were composed of acetate sheets) (47), and constant-depth film fermentor (CDFF; Mark III; University College Cardiff Consultants Ltd.) (50). All systems were run in a once-flow-through configuration. Flow of medium was allowed to occur for at least 3 h prior to inoculation to precondition the system. Flow was stopped, and inoculation was done through the appropriate port using an early-stationary-phase culture (optical density at 470 nm ≈ 2). Flow was resumed after 1 h. In the case of the drip reactor, substrata were sterilized in 75% (vol/vol) ethanol (2 h), rinsed in sterile water, bathed in culture (1 h), and then aseptically transferred to the assembly. In the case of the CDFF, inoculum was fed into the model for 1 h at 0.1 ml · min−1 and then the flow was switched to sterile medium. With the exception of the agar plate model, unless stated, all of the models used were run at room temperature (~20°C) with TSB (one-fifth strength) at a flow rate of 0.1 ml · min−1. All biofilms, apart from the agar plate model, were grown for 7 days prior to analyses. Agar plate biofilms were grown on freshly prepared plates at 37°C for 24 h. One milliliter of the inoculum was swirled over the surface, and excess liquid was then removed.

In the second study, a single system was used (the drip reactor) and medium was altered. The media used were TSB (full, 1/5, and 1/10 strength), LB-Miller (Difco; Becton Dickinson and Company; one-fifth strength), and simple salts medium (pH 5.5 and pH 7.5) (44, 54). All experiments were run at room temperature and a flow rate of 0.1 ml · min−1.

Biofilm samples from outside the laboratory.

Preformed biofilms were also obtained from sources outside the laboratory. These included samples from domestic household water drains (kitchen and bathroom), rotating biological contactors (municipal wastewater treatment facility), a pulp paper factory, water treatment membranes, the glass walls of a freshwater fish aquarium, a water storage tank, and a riverbed (Grand River, Elora, Ontario, Canada). Samples were excised in situ and transported to the lab for embedding and thin sectioning as described below.

Transmission electron microscopy studies.

Samples that were to be assessed by negative staining were prepared as follows. A Formvar- and carbon-coated copper grid (200 mesh; Marivac) was floated, film side down, on 20 μl of sample for 20 s. The grid was removed and the edge gently touched to filter paper (Whatman no. 1; Fisher) so as to wick off excess sample. The grid (sample side) was then washed by being floated on 50 μl of nanopure water, blotted, floated on 10 μl of 2% (wt/vol) uranyl acetate for 10 to 20 s, and blotted dry.

Biofilm samples that were to be thin sectioned were embedded as follows. Each sample was placed in 2.5% glutaraldehyde (vol/vol; in 0.1 M phosphate buffer, pH 7.4; Fisher Scientific) for 0.5 h, rinsed twice with phosphate buffer (0.1 M, 7.4, 5 min), and fixed with osmium tetroxide (1% [wt/vol] in 0.1 M phosphate buffer, pH 7.4; Fisher Scientific) for 0.5 h. Samples were washed twice with nanopure water (5 min) and subjected to an ethanol dehydration series (prepared as volume/volume; Commercial Alcohols Inc.) as follows: 25% (5 min), 50% (5 min), 75% (5 min), 95% (10 min), and 100% (5 min, 5 min, and 10 min). The ethanol was replaced with LR White-ethanol (1:1; London Resin Company, Marivac) for 30 min and then with two changes of 100% LR White over 45 min. The sample was polymerized in fresh LR White (60°C, 1 h) and was ready for thin sectioning. Sections were stained with uranyl acetate and lead citrate.

All samples were examined using a Philips CM10 transmission electron microscope operating at an acceleration voltage of 80 kV under standard operating conditions.

Isolation of matrix and MVs from biofilm.

Matrix was isolated from biofilms grown on agar plates (TSA; diameter, 90 mm; one plate per sample). The biofilms were grown to their equivalent early stationary phase (24 h, determined by dry weight), and biomass was scraped from the surface of the plates and resuspended in sterile 0.9% (wt/vol) NaCl. A homogeneous solution was obtained after vortexing (3 min), and this was centrifuged (12,000 × g, 20 min). The supernatant was retained, and the pellet was resuspended in a volume of saline equivalent to that removed and centrifuged. This was repeated three times, and the pooled supernatant was centrifuged at 12,000 × g (20 min) to pellet whole cells. This was repeated three times, the centrifuged supernatant each time being decanted into to a clean tube. The supernatant at this point was essentially isolated matrix material (see Results below), and from this point forward, MV isolation proceeded as for the supernatant obtained from planktonic cultures (51). After ultracentrifugation, the supernatant was retained for further analyses and the MV pellet was resuspended in 50 mM HEPES (pH 6.8) and frozen (−20°C) for later use.

Isolation of MVs from planktonic populations.

Planktonic cultures were grown in Erlenmeyer flasks using TSB (one-fifth nominal volume of flask). Inoculation (0.5%, vol/vol) was made from an early-stationary-phase culture, and preparations were incubated at 37°C, 125 rpm. Isolation of planktonic MVs was done according to the protocol of Renelli et al. (51).

Quantitative and qualitative characterization of MVs.

Dry weights of samples were obtained by freeze-drying preparations. For dry weight observations, biomass collections were amalgamated in order to obtain sufficient yields.

Diameter values for MVs were determined from micrographs of three independent samples of negatively stained whole mounts, prepared as described previously. Images were archived and analyses were done using the iTEM program (version 5.0; Soft Imaging Systems, Münster, Germany).

CFU were calculated for planktonic and biofilm populations by serial dilution in sterile 0.9% NaCl and triplicate plating of the relevant dilutions on TSA plates. Biofilms were vortexed to obtain a monodisperse suspension before plating. After approximately 20 h of growth, representative plates each having between 30 and 300 colonies were counted and CFU estimates were made. Three independent calculations were performed (n = 9).

3-Deoxy-d-manno-octulosonic acid (Kdo) was evaluated by the periodic acid-thiobarbituric acid method described by Hancock and Poxton (28). Hydrolysis was done for 8 min. Three independent samples were assayed in triplicate (n = 9), and all reagents were purchased from Sigma-Aldrich.

Protein content was quantified using a microbicinchoninic acid protein assay kit (Pierce Bioassay) with bovine serum albumin as the standard. Three independent samples were assayed in triplicate (n = 9).

Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) was used to assess the protein composition of samples and was done following the method described by Kadurugamuwa and Beveridge (34). Forty micrograms of protein was loaded per lane. Broad-range molecular weight markers were purchased from Bio-Rad. Gels were stained overnight with Coomassie blue R-250 (1 g [wt/vol] in 1 liter of methanol-glacial acetic acid-water [5:1:4]) and destained using several changes of methanol-glacial acetic acid-water (1:2:9) before being washed in water to remove all organic solvents. To ensure repeatability of the final composite gel, SDS-PAGE was performed upon duplicated independent experiments, each of which comprised three separate replicates (n = 6). LPS banding profiles were also assessed (20 μg protein per lane) (34). Zymograms for proteolytic enzyme activity (34) (0.15% [wt/vol] gelatin) were performed using samples prepared with or without reducing agent. Gels were archived using a GS-800 densitometer (Bio-Rad) and the Quantity One program (version 4.4.0; Bio-Rad), with the red filter (Coomassie blue) or green filter (silver stain) in place and at a scanning resolution of 63 μm.

Binding of gentamicin by MVs.

Previous studies of gentamicin-induced MV production (34) incubated 8 mg gentamicin/liter culture for 30 min prior to MV harvesting. In order to assess binding of gentamicin specifically to MVs, an aliquot of MVs equivalent in wet weight to that obtained from a planktonic population was incubated with gentamicin (Sigma). This was prepared as a 100-ml MV aliquot equivalent resuspended in 0.2 ml 50 mM HEPES (pH 6.8) to which was added 0.8 mg of gentamicin resuspended in 0.2 ml 50 mM HEPES (pH 6.8). Buffer with no gentamicin served as the negative control. The sample was incubated at room temperature for 30 min, rinsed three times, and concentrated using a Biomax Ultrafree centrifuge filtering system (10-kDa molecular weight cutoff; Millipore). The concentrate was enrobed in an equal volume of Noble agar (4%, wt/vol) and embedded following a less harsh protocol (7) than that outlined earlier so as to minimize alteration of the antigen epitope. Thin sections were immunolabeled (7) using rabbit antigentamicin antiserum (Sigma; 1:100) and goat anti-rabbit immunoglobulin G conjugated to gold (diameter, 10 nm) as the secondary antibody (Sigma; 1:50). Assessment of binding was done using TEM. Less than 1% of bound gold conjugate was seen in the negative control compared to the positive label, and the experiment was repeated in duplicate with observation upon two independent immunolabeled preparations per replicate.

RESULTS AND DISCUSSION

MVs are present within the matrix formed by P. aeruginosa PAO1 biofilms.

Preliminary assessment of the physical presence of MVs was done using the simple yet effective agar plate biofilm model. This model has been widely employed (4, 11, 56, 61, 66), primarily due to its ease of use and generous production of biomass and also because it resembles a soft tissue infection (24). As MVs of P. aeruginosa typically have a diameter of 50 to 150 nm (34), our main means of observation was TEM. In this approach, we used TEM of thin-sectioned biofilms to reveal the internal structure of the biofilm (Fig. (Fig.1a),1a), and in all sections viewed (n = 8; four sections from two independent preparations), MVs were present within the matrix of the biofilm. The distribution of MVs was variable in that there were areas of little MV accumulation and areas that were dense. While thin sections have a thickness of ~60 nm and represent a small sampling of the entire specimen, through the observation of a number of samples it was clear that MVs were consistently seen throughout the entire volume of the biofilm. Mechanical disruption of the biofilms and subsequent whole-mount preparations provided an alternative method to show the presence of the MVs. Here, the MVs were seen associated within aggregates of extracellular material (Fig. (Fig.2a2a).

FIG. 1.
Membrane vesicles in P. aeruginosa PAO1 biofilms. Micrographs of thin sections through embedded P. aeruginosa PAO1 biofilms are shown. The biofilms were grown on a TSA plate (24 h) (a), in a drip reactor (simple salts medium [54], 0.1 ml · min ...
FIG. 2.
Membrane vesicles were present in mechanically disrupted P. aeruginosa PAO1 biofilm and isolated matrix material and were also isolated following a modified MV isolation protocol. Micrographs of negatively stained whole mounts of P. aeruginosa PAO1 biofilm ...

MVs are present within biofilms formed under a variety of conditions.

The next question that needed to be addressed was whether MVs were a common feature within biofilms or whether our observations were a chance occurrence of the system that we had employed. Biofilms of P. aeruginosa PAO1 were grown using different systems (agar plates, flow cells, drip reactors, tubing, and CDFF) and a variety of media. The different systems were chosen to represent some of the conditions under which natural biofilms are formed: saturated (flow cell and silicone tubing), unsaturated (agar plate and drip reactor), aggressive shear (CDFF), fluid flow (drip reactor, silicone tubing, and flow cell), and closed (agar plate) environments. Figure Figure11 shows micrographs of thin sections through these biofilms (Fig. 1a to d).

Again, in contrast with MVs from planktonic bacteria that migrate from the parent cell once liberated, biofilm MVs were found to accrete within the extracellular matrix. This was a consistent feature of all of the different biofilm systems and growth conditions assessed and supported our previous observations. MVs were also located at the substratum-bacterium interface (Fig. (Fig.1b),1b), as a result of either predeposition before cell attachment or production after cell adhesion. In addition, negative stains of effluent from each growth system revealed that MVs were also shed (data not shown). That is, the biofilms not only maintained substantial numbers of MVs within their matrices but also dispersed MVs into the external environment. In this manner, biofilms could effectively serve as both “safe house” factories and depositories of MVs, as well as export depots during processes such as infection or predation and ecological succession. This bears a number of implications in that the traditional boundaries of both cell and intact biofilm become blurred. MVs, as extracellular extensions of cells, may then manifest their effect(s) not only throughout the matrix but also beyond the borders of the biofilm.

Finally, biofilms of Shewanella oneidensis MR-1, Escherichia coli K-12, and an Azotobacter sp., all grown using a drip reactor and one-fifth-strength TSB, were prepared for viewing as thin sections (data not shown). Again, MVs were found to be part of the matrix of these gram-negative bacterial biofilms. This was in agreement with the concept that MV production is a normal function of the physiology of gram-negative bacteria and, in our study, that MVs are a component of the matrix of gram-negative bacterial biofilms.

MVs are found within isolated biofilm matrix material and can be enriched from this fraction.

Biofilms were suspended in saline, and the cells were separated from the matrix by differential centrifugation. Examination of the cell pellet by TEM showed that the majority of the material had been disassociated from the cells, yet ca. 10 to 15% remained bound to cells in the pellet. We did not employ severe chemical or mechanical treatment to fully strip this material from the cells due to concerns about possible cell damage or lysis and subsequent sample contamination. For example, EDTA, a known perturbant of the outer membrane that artificially generates MVs (T. J. Beveridge, unpublished data), has sometimes been employed to increase the yield of isolated extracellular polymeric material (which would be contaminated with abnormal quantities of MVs). TEM of negative stains of our extracted matrix showed the presence of MVs, as well as other particulate structures such as flagella, pyocins, and pili/filamentous phage (10) (Fig. (Fig.2b).2b). Since MVs are blebs from the outer membrane, then Kdo, a marker for LPS, should be present. Indeed, our results were in agreement with this (Table (Table2).2). Previous reports have indicated Kdo to be present within matrix material (22, 23, 66), and we believe that this “matrix LPS” is partly due to the presence of MVs (see below).

TABLE 2.
Kdo values in the various fractions isolated from biofilms

The matrix-associated MVs were then isolated as an enriched preparation by integrating an MV isolation protocol based upon that of Renelli et al. (51). After centrifugation at 150,000 × g, the pellet was enriched with MVs; however, other particulates were also still present—flagella, pili, and pyocins (Fig. (Fig.2c).2c). This was consistent with similar observations of planktonic pellets, since a low-speed centrifugation of the resuspended pellet largely reduced the presence of such contaminating materials from this preparation (51). Kdo values (Table (Table2)2) indicated that this enriched pellet contained a substantial 52% of the LPS present within the matrix material, and dry weight values indicated that this pellet represented 18% (wt/wt) of the matrix material (Table (Table1).1). TEM observations of negative stains of the associated supernatant indicated no MVs to be present, and it is possible that LPS may be associated with other molecules such as polysaccharides, nucleic acids, and proteins which, under the present conditions, did not pellet out. However, MVs clearly represent a major reservoir of LPS within the matrix.

TABLE 1.
Dry weights of biofilm and planktonic fractions

MVs from planktonic and biofilm populations differ both quantitatively and qualitatively.

Upon isolation of the MV-enriched pellets, it was surprising that the volume, color, and consistency of these biofilm MV-enriched pellets contrasted sharply with those obtained from planktonic populations (Fig. (Fig.3).3). The biofilm pellets were consistently larger than the planktonic ones even though they were derived from parent populations of approximately equivalent dry weight and CFU (Table (Table11 and legend to Fig. Fig.3),3), and were of a mid- to dark brown color whereas the planktonic pellets were small and black. The biofilm pellet was more gelatinous than that from planktonic cells (more friable). Negative stains revealed that both preparations had MVs as a major pellet constituent. Together, these observations suggested both qualitative and quantitative differences that required further investigation.

FIG. 3.
MV-enriched ultracentrifuged pellet from planktonic and biofilm populations. MV-enriched ultracentrifuged pellets obtained from CFU-equivalent biofilm populations (right) were of a higher volume and a different color and consistency (more gelatinous) ...

The low-speed centrifugation used to clean contaminating materials from isolated planktonic MVs (51) was implemented for both planktonic and biofilm MVs. In the case of the former, there was an effective reduction in the presence of flagella, pyocins, and pili as determined by TEM of negatively stained samples (Fig. (Fig.4a).4a). Biofilm MVs displayed an anomalous behavior: MVs did sediment, but the vast majority remained in suspension and could be harvested from the supernatant (TEM) (Fig. (Fig.4c).4c). This behavior may be the result of either buoyant density differences or their smaller size (Fig. (Fig.4a4a and and4c;4c; see below). Furthermore, the pellet consisted of an entanglement of flagella, pili, pyocins, and some MVs (Fig. (Fig.4b).4b). The MVs that did pellet tended to have a larger diameter and were more similar in size to the planktonic MVs. Kdo values of the two fractions supported this observation, with more Kdo present in the supernatant than in the pellet (Table (Table2).2). In our hands, this sedimentation behavior was a consistent feature only if the ultracentrifuged sample was sufficiently diluted and resuspended to monodispersity; overloading of the sample resulted in loss of separation. From this point forward, unless otherwise indicated, work with planktonic MVs refers to MV isolations that were pelleted and work with biofilm MVs refers to MVs that remained in suspension. In both instances there was loss of sample in that MVs did not totally migrate to one or the other of the collected fractions with either population.

FIG. 4.
Outcome of low-speed centrifugation on biofilm MVs. MV-enriched ultracentrifuged pellets were resuspended in HEPES buffer and centrifuged at 16,000 × g for 30 min. Planktonic-derived MVs pelleted out (a), whereas the pellet obtained from a biofilm ...

MV yield comparisons were made using the main parameters of dry weight and Kdo. In all instances, under the stated growth conditions, dry weight values indicated that the biofilms exceeded planktonic productivity (Table (Table1).1). Furthermore, given that the biofilm MV wash fraction contained the majority of MVs (see previous paragraph), a revised comparison of the normalized dry weights of the MV-rich fraction from the two populations yields a planktonic/biofilm ratio of 16.38. Kdo values (Table (Table2)2) also indicated a higher yield for biofilm populations, and the ratio of Kdo values (normalized to 1 × 109 CFU) indicated that biofilms produced ca. 120 times more MV-associated Kdo (242 ng versus 2 ng). This sevenfold discrepancy between the weight and Kdo ratios (16.4 versus 121) suggested that biofilm MVs contained more Kdo per unit weight than did their planktonic counterparts. Kdo/protein ratios for the planktonic and biofilm MVs also reflected a similar trend. Planktonic MVs contained 10,650 ng Kdo/g culture protein with a corresponding 15.59 μg Kdo/mg MV protein, whereas biofilm MVs possessed 75,423 ng Kdo/g culture protein with 109.63 μg Kdo/mg MV protein. In both instances, the difference between the respective ratios was sevenfold. SDS-PAGE of Hitchcock-Brown preparations also supported this observation (Fig. (Fig.5a),5a), and biofilm MVs stained more intensely than planktonic MVs when equivalent amounts of protein per sample were loaded. While this sevenfold difference initially appeared odd, a possible explanation may be found within the size distribution data for planktonic and biofilm MV populations (Table (Table3).3). Biofilm MVs were generally smaller, as observed by negative stains and thin sections, which may in part also explain the sedimentation differences noted above. The average MV diameter values obtained for biofilm and planktonic populations were 45 and 86 nm, with corresponding medians of 40 and 85 nm, respectively. This smaller diameter meant that biofilm MVs have an average surface area-to-volume ratio that is approximately twice that of planktonic MVs. This would be reflected in the higher Kdo ratio, but it does not entirely account for the discrepancy in values and supports a genuine difference between the two populations. Our planktonic Kdo values are similar to previously published values (i.e., 66 nmol Kdo/mg protein compared to 41 nmol Kdo/mg protein) (51), yet the values obtained for the biofilm MVs had an astonishing 460 nmol Kdo/mg protein. One possible explanation is that this was a deliberate alteration of the Kdo/protein ratio by the biofilm cells or that the outer membranes from which the MVs are derived are different in planktonic and biofilm populations. Either possibility bears a number of interesting implications. For example, one may consider the reactivity of the LPS molecule and the interactions that it may mediate, the possibility of less protein being packaged into the MV lumen, or the chance that relatively fewer outer membrane proteins occur. The latter may minimize the chances of detection by the host immune system.

FIG. 5.
SDS-PAGE analyses of MVs. SDS-PAGE was used to analyze both planktonic and biofilm MVs. The relative amounts of LPS and protein were compared using silver staining (a), and protein profiles were compared using Coomassie blue staining (b). Finally, proteolytic ...
TABLE 3.
MV diameter distributionsa

Indeed, SDS-PAGE analyses of the two MV-rich populations demonstrated that apart from the quantitative yield differences earlier described, there were dissimilarities other than size and centrifugal sedimentation properties (Fig. (Fig.5b).5b). The planktonic and biofilm MV profiles demonstrated that, although there were proteins in common, there were also a few prominent differences. For example, biofilm MVs have a denser band at around the 55-kDa mark, which could be alkaline protease (see below). In general, however, biofilm MVs had less diversity in the bands present than those from planktonic cells, again supporting the notion that a more limited selection of proteins is being secreted as part of the biofilm MV complement. Future studies into the nature of the different proteins would shed light onto this and perhaps reveal some commonality in those proteins that are present.

Investigations into the functionality of biofilm MVs.

Due to the large economic expense that MV production entails to the cell, it has long been argued (and demonstrated) that they are not nonsensically produced. Among the diverse functions ascribed to planktonic MVs are their roles as a novel secretory pathway (37, 60), the delivery of virulence factors (20, 30, 31, 34), cell-to-cell signals (42), cell aggregation (19, 25, 35), metal immobilization and redox processes affecting minerals, inactivation of antimicrobials by enzymatic degradation (e.g., β-lactamases) (13) or by binding (26), the selection and destruction of non-self cells (40), immune-modulating substances (1, 45, 46), and (somewhat controversially) the transfer of genetic material (17, 39, 68). All of these processes do occur within the scope of a biofilm, and there is no apparent reason why MVs should not participate in them. Furthermore, given that the MVs are shed, these processes may be relayed to actively have an effect outside of the confines of the biofilm, even at a good distance from it.

Planktonic MVs have been reported to contain potent hydrolytic enzymes such as proteases and peptidoglycan hydrolases (34, 40). As the SDS-PAGE analyses (Fig. (Fig.5b)5b) had suggested the possibility of a protease, we performed zymogram assays (34). Samples were prepared both in the presence and in the absence of a reducing agent. This was essential as certain enzymes lose their functionality or give an enhanced response in the presence of a reducing agent (41). Proteolytic activity was found associated with both the planktonic and biofilm MVs (Fig. (Fig.5c),5c), under both conditions. However, in both instances using the same sample concentrations, biofilm MVs demonstrated greater proteolytic activity, as denoted by the larger zones of clearing. Under nonreducing conditions, four bands were seen: a very diffuse band at the 20-kDa region, a band at approximately 55 kDa, and two bands running at 80 to 90 kDa. It was noted that the Rf values of some bands differed from those run on a gel without gelatin. Under reducing conditions, only one prominent band was present at approximately 55 kDa. This band is consistent with alkaline protease, as is its behavior under reducing conditions (41). It is possible that the diffuse 20-kDa-region band seen under nonreducing conditions was PrpL (27 kDa) (53). However, the evidence remains that biofilm MVs contained more proteolytic activity than those from planktonic cells, an important consideration when thinking of the roles that MVs may play in pathogenesis, release of nutrients, and perhaps surface modification. Furthermore, there are reports of proteases within the matrices of biofilms (53), and clearly, a proportion of these reside in MVs.

In the second study, we assessed the ability of MVs to bind antibiotics, in particular gentamicin. The binding of this cationic aminoglycoside is well studied (33) and a logical starting point. Both planktonic and biofilm MVs were incubated with gentamicin, and binding was assessed by immunolabeling and TEM (Fig. (Fig.6).6). The micrographs clearly demonstrate that biofilm MVs did bind gentamicin, and the label revealed the aminoglycoside to be located at the outer and inner faces of the membrane, as well as within the lumen. Planktonic MVs were also shown to interact with gentamicin (data not shown). Clearly, since gentamicin was attached to all regions of the MVs, the antibiotic must have penetrated entirely through the MV, indicating strong or even disruptive bilayer interaction. This is not surprising since highly cationic substances, such as aminoglycoside antibiotics, can displace essential metal cations and disrupt lipid-packing order (33). Not all MVs showed labeling, but this is expected and was seen in other studies (34). This could be due to a number of factors, e.g., diminished antibody-antigen affinity due either to the antibody itself (the manufacturer's sheet states that binding is up to 40% of gentamicin present for a radioimmunoassay) or antigen alteration during the embedding protocol, or it could be that the bound gentamicin was simply not accessible in the thin section. Indeed, in our experience the consistent labeling of these small vesicles was remarkably efficient considering their size and likely exposure in thin section (7). Yet, it remains that the MVs did bind exogenous gentamicin, and given the surface chemistry of MVs, the vesicles could also bind other extraneous compounds. As a general property of biofilm MVs, they could act as decoys or “sponges” to reduce inimical agents within biofilms before they affect cells. Since MVs are released from biofilms, the concentration of such agents within the biofilm would be reduced and MVs dispersed from a biofilm could serve to bind agent prior to contact with the biofilm. This could be another way in which biofilms are protected from antimicrobials. If so, it represents a simple but effective mechanism and a route that we hope to explore in greater detail.

FIG. 6.
Binding of gentamicin by MVs. Shown are micrographs of biofilm MVs that had been incubated with gentamicin prior to embedding and thin sectioning, probed with a primary antibody specific for gentamicin, and developed with a secondary gold-conjugated antibody. ...

MVs are also found within naturally occurring biofilms.

Laboratory-grown biofilms are very different from natural biofilms. The former are often monocultures subjected to a constant and reasonable flow of nutrients, whereas the latter are a complex mixture of cultures that are subjected to a variable and often harsh set of environmental conditions. It was therefore necessary to sample natural biofilms to ensure that MVs were a natural trait of these systems. A number of different environmental conditions were sampled. These included domestic water drains, sewage and water treatment plants, pulp and paper manufacturers, freshwater fish aquariums, water storage tanks, and riverbeds. MVs were seen in all biofilms where gram-negative bacteria were found. There was a more discontinuous distribution of MVs than that seen in any of the PAO1 biofilms. This was presumably related to the distribution of gram-negative bacteria present since MVs were shed and found close to these regions (Fig. (Fig.7),7), although other factors such as environmental conditions could play a role (30, 36, 52). Additionally, environmental biofilms often incorporated more extraneous material (e.g., large plant fibers, mineral grains, large protists, etc.), effectively adding to the bulk of the biofilm. Our laboratory has extensive experience in sampling microbiota growing as biofilms or flocs (especially those of marine “whiting” events) from a number of dissimilar natural environments. We have never encountered a natural setting where gram-negative bacteria were present that did not contain MVs.

FIG.7.FIG.7.
Membrane vesicles in biofilms from outside the lab. Shown are micrographs of thin sections through conventionally embedded biofilms obtained from a domestic bathroom drain (a) and a water treatment membrane (b). Note the presence of different cell types. ...

Potential of biofilm MVs.

The overall goal of our study was to establish MVs as a bona fide component of the matrix of gram-negative bacterial biofilms. Through our TEM studies, we found that MVs were a definite and consistent component of the matrix of gram-negative bacterial biofilms. Furthermore, they were shed from the biofilms, disseminating into the external environment where they could serve as a source of extracellular activity both in and ex biofilm. Under defined conditions, we found that these exquisite structures could account for up to 16.8% (wt/wt) of the matrix material and more than 50% of the Kdo found within the matrix material. The presence of these LPS-bearing structures firmly places LPS as a matrix component and not a contaminant or procedural artifact. Additionally, while Kdo values have been used to show that LPS is a minor-percentage matrix component, Kdo represents only 2 to 6% of the LPS (wt/wt) of P. aeruginosa PAO1, a fact that is often overlooked. These biofilm MVs were also found to substantially differ from their planktonic counterparts in terms of their physical dimensions and properties, as well as in their chemistry. It was impossible for us to probe MVs for all potential functions, but we have certainly indicated a few—the packaging of virulence factors such as proteases and the binding of antibiotics.

In some of the samples, MVs were located at the substratum-biofilm interface and either were deposited prior to biofilm development or were produced by the biofilm, i.e., cells can alter the surface properties of the substratum either pre- or post-biofilm formation. Planktonic MVs derived from Bacteroides gingivalis adhere to hydroxyapatite and facilitate the attachment of Streptococcus sanguis (55). Other studies have indicated that MVs can also mediate aggregation of cells (19, 35, 43). Can cells then use MVs to influence early-stage biofilm processes such as adhesion? It is important to recognize that biofilms form on an enormous variety of substrata that range from inert mineral faces to high-carbon films (e.g., cellulose and chitin) to soft tissue surfaces (plant, mammalian, etc.); MVs could be actively or passively altering substratum surface properties. Alternatively, the MVs could simply be entrapped between the substratum and the producing cells.

Substratum-associated MVs, however, account for only a fraction of the total. In considering the size, frequency, and chemical nature of MVs, these occupied a substantial volume of the biofilm. It is also important to remember that MVs are characteristic of the producing cell and its phenotype, possessing serotype-specific LPS and outer membrane proteins, which provide a unique surface chemistry for environmental interactions. Available surface molecules or intrinsic reducing activity could affect and be affected by the redox and pH of the local microenvironment surrounding groups of cells within the biofilm. These very same interactions might play a role in stabilizing polymers, ions, and other components of the matrix, all contributing to the properties of a microenvironment. Close to the producer cells, through small-scale interactions, they could have an impact by providing sorbative or inactivating power on extraneous agents, thereby protecting cells. Additionally, MVs could also be capable of interacting with polymers within the biofilm, e.g., DNA and polysaccharide, influencing their ability for entanglement. This must alter the intrinsic properties of the polymers through electrostatic or hydrophobic interaction and binding. Indeed, DNA, a substantial ingredient of the matrix of P. aeruginosa biofilms (3, 63), is a proven constituent of MVs and can be associated with the lumen or the membrane surface (34, 51). Accordingly, some of this matrix DNA could actually be MV DNA. There could also be strong associations between exogenous matrix DNA and MVs as well as other biofilm particulates (58). It is even possible that MVs could contain enzymes capable of altering polymers (e.g., nucleases, polysaccharases, and epimerases). All particulates and their interactions with the matrix must have a strong impact on the rheological properties of biofilms.

Apart from the possibility of physical interaction with extracellular polymers, MVs have been shown to contain active periplasmic components, e.g., enzymes and toxins. For example, certain strains of P. aeruginosa produce and package β-lactamase into MVs, which can then degrade β-lactam antibiotics (13). It is interesting that biofilms exposed to β-lactams have an increased synthesis of β-lactamase (5, 6). In addition, Porphyromonas gingivalis produces MVs that bind and sequester chlorhexidine (26). It is then possible that biofilm cells can be induced to shed MVs with active properties that would neutralize inimical agents designed to attack and destroy biofilms.

Finally, biofilm MVs appear to be independent, extracellular extensions of the cell, which broaden the traditional boundaries of the cell. These boundaries include regions within the biofilm itself and beyond its confines. In the latter context, we could imagine a biofilm being a location from which MVs are liberated to manifest certain properties in the broad-scale environment. These could be intrinsic chemical properties (e.g., those affecting geochemical conditions within a geological horizon) or active components (e.g., virulence factors) to promote a desired bioeffect. Biofilms would be much more long-lasting durable depots for these bioactive MV particles than planktonic cells and would be a continual source of such environmentally altering substances. Since many of the active ingredients are encapsulated within the MVs, they would be better protected from antagonistic external factors. Furthermore, since MVs mimic the bacterial cell surface, they would have strong adhesive properties or (even) specific adhesions for attachment (e.g., on certain tissue types).

In summary, there is an increased awareness of biofilms and their abundance in nature. Hardly a month goes by without new reports on the properties of biofilms, yet few have concentrated on the particulate properties of the biofilm matrix. While our nascent understanding of the matrices of biofilms led to the belief that these consisted primarily of exopolysaccharides, this study, along with many others, consolidates this as an erroneous perception. The matrix is a complex amalgam, comprised of polymers and macromolecules, as well as particulate structures such as discarded pili and flagella. Here, in our report, we emphasize MVs as being among the most important because of their intrinsic surface properties and active constituents. We hope this will encourage further studies of these fascinating nanoparticles.

Acknowledgments

This work was made possible by an NSERC-Discovery grant to T.J.B. for general operating funds and by CBDN-NCE and AFMnet-NCE grants to T.J.B., which helped cover salaries. The TEM was performed in the NSERC Guelph Regional Integrated Imaging Facility (GRIIF), which is partially funded by an NSERC-MFA grant to T.J.B.

We thank Corinne Michaud for preliminary work that led to the development of this study, Dianne Moyles and Robert Harris for expert guidance with the TEM-related aspects of this study, and Anuradha Saxena for technical assistance.

REFERENCES

1. Aase, A., L. M. Næss, R. H. Sandin, T. K. Herstad, F. Oftung, J. Holst, I. L. Haugen, E. A. Høiby, and T. E. Michaelsen. 2003. Comparison of functional immune responses in humans after intranasal and intramuscular immunisations with outer membrane vesicle vaccines against group B meningococcal disease. Vaccine 21:2042-2051. [PubMed]
2. Allan, N. D., C. Kooi, P. A. Sokol, and T. J. Beveridge. 2003. Putative virulence factors are released in association with membrane vesicles from Burkholderia cepacia. Can. J. Microbiol. 49:613-624. [PubMed]
3. Allesen-Holm, M., K. B. Barken, L. Yang, M. Klausen, J. S. Webb, S. Kjelleberg, S. Molin, M. Givskov, and T. Tolker-Nielsen. 2006. A characterization of DNA release in Pseudomonas aeruginosa cultures and biofilms. Mol. Microbiol. 59:1114-1128. [PubMed]
4. Auerbach, I. D., C. Sorensen, H. G. Hansma, and P. A. Holden. 2000. Physical morphology and surface properties of unsaturated Pseudomonas putida biofilms. J. Bacteriol. 182:3809-3815. [PMC free article] [PubMed]
5. Bagge, N., M. Hentzer, J. B. Andersen, O. Ciofu, M. Givskov, and N. Høiby. 2004. Dynamics and spatial distribution of β-lactamase expression in Pseudomonas aeruginosa biofilms. Antimicrob. Agents Chemother. 48:1168-1174. [PMC free article] [PubMed]
6. Bagge, N., M. Schuster, M. Hentzer, O. Ciofu, M. Givskov, E. P. Greenberg, and N. Høiby. 2004. Pseudomonas aeruginosa biofilms exposed to imipenem exhibit changes in global gene expression and β-lactamase and alginate production. Antimicrob. Agents Chemother. 48:1175-1187. [PMC free article] [PubMed]
7. Beveridge, T. J., T. J. Popkin, and R. M. Cole. 1994. Electron microscopy, p. 42-71. P. Gerhardt, R. G. E. Murray, W. A. Wood, and N. R. Krieg (ed.), Methods for general and molecular microbiology. ASM Press, Washington, D.C.
8. Beveridge, T. J., S. A. Makin, J. L. Kadurugamuwa, and Z. Li. 1997. Interactions between biofilms and the environment. FEMS Microbiol. Rev. 20:291-303. [PubMed]
9. Beveridge, T. J. 1999. Structures of gram-negative cell walls and their derived membrane vesicles. J. Bacteriol. 181:4725-4733. [PMC free article] [PubMed]
10. Bradley, D. E. 1973. The adsorption of the Pseudomonas aeruginosa filamentous bacteriophage Pf to its host. Can. J. Microbiol. 19:623-631. [PubMed]
11. Branda, S. S., Å. Vik, L. Friedman, and R. Kolter. 2005. Biofilms: the matrix revisited. Trends Microbiol. 13:20-26. [PubMed]
12. Caldwell, D. E., D. R. Korber, and J. R. Lawrence. 1992. Confocal laser microscopy and digital image analysis in microbial ecology. Adv. Microb. Ecol. 12:1-67.
13. Ciofu, O., T. J. Beveridge, J. L. Kadurugamuwa, J. Walther-Rasmussen, and N. Høiby. 2000. Chromosomal β-lactamase is packaged into membrane vesicles and secreted from Pseudomonas aeruginosa. J. Antimicrob. Chemother. 45:9-13. [PubMed]
14. Costerton, J. W., G. G. Geesey, and K.-J. Cheng. 1978. How bacteria stick. Sci. Am. 238:86-95. [PubMed]
15. Costerton, J. W., I. T. Irvin, and K.-J. Cheng. 1981. The bacterial glycocalyx in nature and disease. Annu. Rev. Microbiol. 35:299-324. [PubMed]
16. deBeer, D., and P. Stoodley. 1994. Effects of biofilm structure on oxygen distribution and mass transport. Biotechnol. Bioeng. 43:1131-1138. [PubMed]
17. Dorward, D. W., C. F. Garon, and R. C. Judd. 1989. Export and intracellular transfer of DNA via membrane blebs of Neisseria gonorrhoeae. J. Bacteriol. 171:2499-2505. [PMC free article] [PubMed]
18. Dunne, W. M., Jr., F. L. A. Buckmire, and V. M. Kushnaryov. 1982. Comparative ultrastructure of a mucoid strain of Pseudomonas aeruginosa isolated from cystic fibrosis patient and its spontaneous non-mucoid mutant. Microbios 34:197-212. [PubMed]
19. Ellen, R. P., and D. A. Grove. 1989. Bacteroides gingivalis vesicles bind to and aggregate Actinomyces viscosus. Infect. Immun. 57:1618-1620. [PMC free article] [PubMed]
20. Fiocca, R., V. Necchi, P. Sommi, V. Ricci, J. Telford, T. L. Cover, and E. Solcia. 1999. Release of Helicobacter pylori vacuolating cytotoxin by both a specific secretory pathway and budding of outer membrane vesicles. Uptake of released toxin and vesicles by gastric epithelium. J. Pathol. 188:220-226. [PubMed]
21. Forsberg, C. W., T. J. Beveridge, and A. Hellstrom. 1981. Cellulase and xylanase release from Bacteroides succinogenes and its importance in the rumen environment. Appl. Environ. Microbiol. 42:886-896. [PMC free article] [PubMed]
22. Friedman, L., and R. Kolter. 2004. Genes involved in matrix formation in Pseudomonas aeruginosa PA14 biofilms. Mol. Microbiol. 51:675-690. [PubMed]
23. Friedman, L., and R. Kolter. 2004. Two genetic loci produce distinct carbohydrate-rich structural components of the Pseudomonas aeruginosa matrix. J. Bacteriol. 186:4457-4465. [PMC free article] [PubMed]
24. Gilbert, P., and D. G. Allison. 1993. Laboratory methods for biofilm production. Soc. Appl. Bacteriol. Tech. Ser. 30:29-49.
25. Grenier, D., and D. Mayrand. 1987. Functional characterization of extracellular vesicles produced by Bacteroides gingivalis. Infect. Immun. 55:111-117. [PMC free article] [PubMed]
26. Grenier, D., J. Bertrand, and D. Mayrand. 1995. Porphyromonas gingivalis outer membrane vesicles promote bacterial resistance to chlorhexidine. Oral Microbiol. Immunol. 10:319-320. [PubMed]
27. Halhoul, N., and J. R. Colvin. 1975. The ultrastructure of bacterial plaque attached to the gingiva of man. Arch. Oral Biol. 20:115-118. [PubMed]
28. Hancock, I., and I. Poxton. 1988. Appendix 1, p. 273. In I. Hancock and I. Poxton (ed.), Bacterial cell surface techniques. John Wiley & Sons, Chichester, United Kingdom.
29. Hentzer, M., G. M. Teitzel, G. J. Balzer, A. Heydorn, S. Molin, M. Givskov, and M. R. Parsek. 2001. Alginate overproduction affects Pseudomonas aeruginosa biofilm structure and function. J. Bacteriol. 183:5395-5401. [PMC free article] [PubMed]
30. Horstman, A. L., and M. J. Kuehn. 2000. Enterotoxigenic Escherichia coli secretes active heat-labile via outer membrane vesicles. J. Biol. Chem. 275:12489-12496. [PubMed]
31. Horstman, A. L., and M. J. Kuehn. 2002. Bacterial surface association of heat-labile enterotoxin through lipopolysaccharide after secretion via the general secretory pathway. J. Biol. Chem. 277:32538-32545. [PubMed]
32. Hunter, R. C., and T. J. Beveridge. 2005. High-resolution visualization of Pseudomonas aeruginosa PAO1 biofilms by freeze-substitution transmission electron microscopy. J. Bacteriol. 187:6719-6730. [PMC free article] [PubMed]
33. Kadurugamuwa, J. L., A. J. Clarke, and T. J. Beveridge. 1993. Surface action of gentamicin on Pseudomonas aeruginosa. J. Bacteriol. 175:5798-5805. [PMC free article] [PubMed]
34. Kadurugamuwa, J. L., and T. J. Beveridge. 1995. Virulence factors are released from Pseudomonas aeruginosa in association with membrane vesicles during normal growth and exposure to gentamicin: a novel mechanism of enzyme secretion. J. Bacteriol. 177:3998-4008. [PMC free article] [PubMed]
35. Kamaguchi, A., K. Nakayama, S. Ichiyama, R. Nakamura, T. Watanabe, M. Ohta, H. Baba, and T. Ohyama. 2003. Effect of Porphyromonas gingivalis vesicles on Staphylococcus aureus aggregation to oral microorganisms. Curr. Microbiol. 47:485-491. [PubMed]
36. Keenan, J. I., and R. A. Allardyce. 2000. Iron influences the expression of Helicobacter pylori outer membrane vesicle-associated virulence factors. Eur. J. Gastroenterol. Hepatol. 12:1267-1273. [PubMed]
37. Kesty, N. C., K. M. Mason, M. Reedy, S. E. Miller, and M. J. Kuehn. 2004. Enterotoxigenic Escherichia coli vesicles target toxin delivery into mammalian cells. EMBO J. 23:4538-4549. [PMC free article] [PubMed]
38. Klausen, M., A. Heydorn, P. Ragas, L. Lambertsen, A. A. Aaes-Jørgensen, S. Molin, and T. Tolker-Nielsen. 2003. Biofilm formation by Pseudomonas aeruginosa wild type, flagella and type IV pili mutants. Mol. Microbiol. 48:1511-1524. [PubMed]
39. Kolling, G. L., and K. R. Matthews. 1999. Export of virulence genes and Shiga toxin by membrane vesicles of Escherichia coli O157:H7. Appl. Environ. Microbiol. 65:1843-1848. [PMC free article] [PubMed]
40. Li, Z., A. J. Clarke, and T. J. Beveridge. 1998. Gram-negative bacteria produce membrane vesicles which are capable of killing other bacteria. J. Bacteriol. 180:5478-5483. [PMC free article] [PubMed]
41. Lomholt, J. A., K. Poulsen, and M. Kilian. 2001. Epidemic population structure of Pseudomonas aeruginosa: evidence for a clone that is pathogenic to the eye and that has a distinct combination of virulence factors. Infect. Immun. 69:6284-6295. [PMC free article] [PubMed]
42. Mashburn, L. M., and M. Whiteley. 2005. Membrane vesicles traffic signals and facilitate group activities in a prokaryote. Nature 437:422-425. [PubMed]
43. Mayrand, D., and D. Grenier. 1989. Biological activities of outer membrane vesicles. Can. J. Microbiol. 35:607-613. [PubMed]
44. McGrath, J. W., S. Cleary, A. Mullan, and J. P. Quinn. 2001. Acid-stimulated phosphate uptake by activated sludge microorganisms under aerobic laboratory conditions. Water Res. 35:4317-4322. [PubMed]
45. Mirlashari, M. R., E. A. Høiby, J. Holst, and T. Lyberg. 2001. Outer membrane vesicles from Neisseria meningitidis: effects on cytokine production in human whole blood. Cytokine 13:91-97. [PubMed]
46. Moe, G. R., P. Zuno-Mitchell, S. N. Hammond, and D. M. Granoff. 2002. Sequential immunization with vesicles prepared from heterologous Neisseria meningitidis strains elicits broadly protective serum antibodies to group B strains. Infect. Immun. 70:6021-6031. [PMC free article] [PubMed]
47. Nguyen, T. T. 2005. M.Sc. thesis. University of Guelph, Guelph, Ontario, Canada.
48. Nivens, D. E., D. E. Ohman, J. Williams, and M. J. Franklin. 2001. Role of alginate and its O acetylation in formation of Pseudomonas aeruginosa microcolonies and biofilms. J. Bacteriol. 183:1047-1057. [PMC free article] [PubMed]
49. Parsek, M. R., and E. P. Greenberg. 1999. Quorum sensing signals in development of Pseudomonas aeruginosa biofilms. Methods Enzymol. 310:43-55. [PubMed]
50. Peters, A. C., and J. W. T. Wimpenny. 1988. A constant-depth laboratory model film fermenter. Biotechnol. Bioeng. 32:263-270. [PubMed]
51. Renelli, M., V. Matias, R. Lo, and T. J. Beveridge. 2004. DNA-containing membrane vesicles of Pseudomonas aeruginosa PAO1 and their genetic information potential. Microbiology 150:2161-2169. [PubMed]
52. Sabra, W., H. Lünsdorf, and A.-P. Zeng. 2003. Alterations in the formation of lipopolysaccharide and membrane vesicles on the surface of Pseudomonas aeruginosa PAO1 under oxygen stress conditions. Microbiology 149:2789-2795. [PubMed]
53. Sarkisova, S., M. A. Patrauchan, D. Berglund, D. E. Nivens, and M. J. Franklin. 2005. Calcium-induced virulence factors associated with the extracellular matrix of mucoid Pseudomonas aeruginosa biofilms. J. Bacteriol. 187:4327-4337. [PMC free article] [PubMed]
54. Schooling, S. R., U. K. Charaf, D. G. Allison, and P. Gilbert. 2004. A role for rhamnolipid in biofilm dispersion. Biofilms 1:91-99.
55. Singh, U., D. Grenier, and B. C. McBride. 1989. Bacteroides gingivalis vesicles mediate attachment of streptococci to serum-coated hydroxyapatite. Oral Microbiol. Immunol. 4:199-203. [PubMed]
56. Strathmann, M., J. Wingender, and H.-C. Flemming. 2002. Application of fluorescently labelled lectins for the visualization and biochemical characterization of polysaccharides in biofilms of Pseudomonas aeruginosa. J. Microbiol. Methods 50:237-248. [PubMed]
57. Vallet, I., J. W. Olson, S. Lory, A. Lazdunski, and A. Filloux. 2001. The chaperone/usher pathways of Pseudomonas aeruginosa: identification of fimbrial gene clusters (cup) and their involvement in biofilm formation. Proc. Natl. Acad. Sci. USA 98:6911-6916. [PMC free article] [PubMed]
58. van Schaik, E. J., C. L. Giltner, G. F. Audette, D. W. Keizer, D. L. Bautista, C. M. Slupsky, B. D. Sykes, and R. T. Irvin. 2005. DNA binding: a novel function of Pseudomonas aeruginosa type IV pili. J. Bacteriol. 187:1455-1464. [PMC free article] [PubMed]
59. Vroom, J. M., K. J. DeGraw, H. C. Gerritsen, D. J. Bradshaw, and P. D. Marsh. 1999. Depth penetration and detection of pH gradients in biofilms by two-photon excitation microscopy. Appl. Environ. Microbiol. 65:3502-3511. [PMC free article] [PubMed]
60. Wai, S. N., B. Lindmark, T. Söderblom, A. Takade, M. Westermark, J. Oscarsson, J. Jass, A. Richter-Dahlfors, Y. Mizunoe, and B. E. Uhlin. 2003. Vesicle-mediated export and assembly of pore-forming oligomers of the enterobacterial ClyA cytotoxin. Cell 115:25-35. [PubMed]
61. Walters, M. C., III, F. Roe, A. Bugnicourt, M. J. Franklin, and P. S. Stewart. 2003. Contributions of antibiotic penetration, oxygen limitation, and low metabolic activity to tolerance of Pseudomonas aeruginosa biofilms to ciprofloxacin and tobramycin. Antimicrob. Agents Chemother. 47:317-323. [PMC free article] [PubMed]
62. Webb, J. S., M. Lau, and S. Kjelleberg. 2004. Bacteriophage and phenotypic variation in Pseudomonas aeruginosa biofilm development. J. Bacteriol. 186:8066-8073. [PMC free article] [PubMed]
63. Whitchurch, C. B., T. Tolker-Nielsen, P. C. Ragas, and J. S. Mattick. 2002. Extracellular DNA required for bacterial biofilm formation. Science 295:1487. [PubMed]
64. Wingender, J., T. R. Neu, and H.-C. Flemming. 1999. What are bacterial extracellular polymeric substances?, p. 1-19. In J. Wingender, T. R. Neu, and H.-C. Flemming (ed.), Microbial extracellular polymeric substances—characterization, structure and function. Springer-Verlag, Berlin, Germany.
65. Wolfaardt, G. M., J. R. Lawrence, and D. R. Korber. 1999. Function of EPS, p. 171-200. In J. Wingender, T. R. Neu, and H.-C. Flemming (ed.), Microbial extracellular polymeric substances—characterization, structure and function. Springer-Verlag, Berlin, Germany.
66. Wozniak, D. J., T. J. O. Wyckoff, M. Starkey, R. Keyser, P. Azadi, G. A. O'Toole, and M. R. Parsek. 2003. Alginate is not a significant component of the extracellular polysaccharide matrix of PA14 and PAO1 Pseudomonas aeruginosa biofilms. Proc. Natl. Acad. Sci. USA 100:7907-7912. [PMC free article] [PubMed]
67. Xu, K. D., P. S. Stewart, F. Xia, C.-T. Huang, and G. A. McFeters. 1998. Spatial physiological heterogeneity in Pseudomonas aeruginosa biofilm is determined by oxygen availability. Appl. Environ. Microbiol. 64:4035-4039. [PMC free article] [PubMed]
68. Yaron, S., G. L. Kolling, L. Simon, and K. R. Matthews. 2000. Vesicle-mediated transfer of virulence genes from Escherichia coli O157:H7 to other enteric bacteria. Appl. Environ. Microbiol. 66:4414-4420. [PMC free article] [PubMed]

Articles from Journal of Bacteriology are provided here courtesy of American Society for Microbiology (ASM)
PubReader format: click here to try

Formats:

Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...

Links

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...