JOURNAL OF BACTERIOLOGY, Apr. 2009, p. 2077–2082 Vol. 191, No. 70021-9193/09/$08.00�0 doi:10.1128/JB.01333-08Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Three-Dimensional Macromolecular Organization of CryofixedMyxococcus xanthus Biofilms as Revealed by Electron

Microscopic Tomography�†Hildur Palsdottir,1 Jonathan P. Remis,1 Christoph Schaudinn,2 Eileen O’Toole,3 Renate Lux,4

Wenyuan Shi,4 Kent L. McDonald,5 J. William Costerton,2 and Manfred Auer1*Lawrence Berkeley National Laboratory, 1 Cyclotron Road, Berkeley, California 947201; Center for Biofilms, School of Dentistry,

University of Southern California, 925 West 34th Street, Los Angeles California 900892; Boulder Laboratory for 3D ElectronMicroscopy of Cells, Dept. MCD Biology, University of Colorado, Boulder, Colorado 803093; UCLA School of Dentistry,

10833 Le Conte Avenue, Los, Angeles, California 90095-16684; and Electron Microscope Laboratory,University of California, Berkeley, California5

Received 23 September 2008/Accepted 12 January 2009

Despite the fact that most bacteria grow in biofilms in natural and pathogenic ecosystems, very little isknown about the ultrastructure of their component cells or about the details of their community architecture.We used high-pressure freezing and freeze-substitution to minimize the artifacts of chemical fixation, sampleaggregation, and sample extraction. As a further innovation we have, for the first time in biofilm research, usedelectron tomography and three-dimensional (3D) visualization to better resolve the macromolecular 3Dultrastructure of a biofilm. This combination of superb specimen preparation and greatly improved resolutionin the z axis has opened a window in studies of Myxococcus xanthus cell ultrastructure and biofilm communityarchitecture. New structural information on the chromatin body, cytoplasmic organization, membrane appo-sition between adjacent cells, and structure and distribution of pili and vesicles in the biofilm matrix ispresented.

Bacteria are usually found concentrated at solid-liquid in-terfaces, where they colonize natural and man-made surfacesin community-like structures termed bacterial biofilms (6).Bacteria in biofilms exhibit protein expression patterns differ-ent from those of planktonic bacteria (41). Within a biofilm,bacteria are typically embedded in an extracellular polymericsubstance (EPS) matrix (7, 17, 43) that protects the microbialcommunity members from desiccation, from phagocytosis,and, in the case of human pathogens, from the host immunesystem (8).

While a variety of biofilms have been studied in depth byoptical imaging approaches, only a few have been faithfullypreserved and visualized at high resolution by transmissionelectron microscopy (3, 22, 25, 26, 45, 48, 49). A recent ultra-structural study of Pseudomonas aeruginosa biofilms shows thathigh-pressure freezing/freeze-substitution results in superior pres-ervation (22). Unlike conventional sample preparation, millisec-ond cryofixation and low-temperature dehydration protocols min-imize artifacts from chemical fixation, extraction, and aggregation(15, 16, 19, 20, 22).

Conventional two-dimensional (2D) imaging provides x,ypositional information but cannot resolve features along the zdirection. Conventional 2D projection imaging of thin (�70- to100-nm) sections results in the superposition of �5 to 20 layers

of protein along the electron path, obscuring molecular details.True 3D visualization requires electron tomographic data ac-quisition and 3D reconstruction, upon which both intracellularand extracellular features are resolved. In this way, cellularorganelles, molecular machines, and macromolecular com-plexes can be visualized in their native microenvironment. Todate, electron tomographic 3D reconstruction of a bacterialbiofilm has not been reported.

Myxococcus xanthus is a well-studied gram-negative soilscavenger and predator (21), and it serves as a model systemfor biofilm formation. M. xanthus moves over surfaces either byadventurous (A) motility or social (S) motility (for a review,see reference 18). It forms spore-bearing fruiting bodies uponstarvation, in a process that requires the coordinated move-ment of many thousands of cells. S motility relies on retractionof type IV pili (T4P) upon interaction with amine-containingpolysaccharides in the extracellular matrix (11, 27, 38, 46). T4Pare also required for cell surface adhesion and “clumping.” Itis unclear how prominent T4P are in stable microbial commu-nities and what their possible role might be. Also, it is unknownhow bacterial community members interact with one anotherand what mechanisms they employ for effective communica-tion and signal and/or material transfer in such dense commu-nities.

Here, we report faithful preservation of M. xanthus biofilmsand the first macromolecular resolution-based insights intotheir 3D architecture, obtained by electron tomographic imag-ing of rapidly frozen, freeze-substituted, and resin-embeddedbiofilm sections. The tomograms reveal the 3D organization ofthe bacterial chromosome, direct cell-cell and cell-substrateinteractions, the presence of extracellular filaments (including

* Corresponding author. Mailing address: Lawrence Berkeley Na-tional Laboratory, 1 Cyclotron Road, Berkeley, CA 94720. Phone:(510) 486-7702. Fax: (510) 486-6488. E-mail: MAuer@lbl.gov.

† This paper is dedicated to the memory of Terry Beveridge, whoserecent and untimely death robbed this field of one of its most percep-tive and competent researchers.

� Published ahead of print on 23 January 2009.

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but not restricted to T4P), and abundant extracellular vesiclesthat, interestingly, we found to be tethered to each other andto the bacterial outer membrane.

MATERIALS AND METHODS

Ten milliliters of CYE broth, consisting of 5 mM MOPS (morpholinepropane-sulfonic acid) (pH 7.6), 2 mM MgSO4, 0.5% (wt/wt) Bacto Casitone, and 0.25%(wt/wt) Bacto yeast extract (5), was inoculated with Myxococcus xanthus DK1622(wild type) and grown for 24 h at 32°C and 100 rpm. Four milliliters of thisculture was used to inoculate 200 ml of CYE broth, which was incubated for 18 hat 32°C at 100 rpm. The culture was concentrated to a final optical density at 600nm of 10. Cellulose microdialysis hollow fibers (Spectra/Por; Spectrumlabs, CA)were cut into 1-mm-long pieces, autoclaved together with 1- by 1-mm2 squares ofIsopore membrane filters (HTTP, 0.4 �m; Millipore, MA), and subsequentlyplaced on CYE agar. Ten microliters of concentrated culture (optical density at600 nm of 10) was placed on top of each Spectra/Por fiber/Isopore filter orIsopore membranes and incubated for 6 h at 32°C followed by 20 h at roomtemperature. In the case of cellulose tubes, biofilms were grown by placing the�200-�m-thick cellulose microdialysis tube with the Myxococcus culture on topof a moist agar plate prior to overnight shipment. Using these culturing condi-tions, the biofilms growing on Isopore filters were noticeably thicker, typically 4to 10 cell layers thick, whereas the biofilms grown on cellulose tubes were thin,sometimes as thin as 2 cell layers.

High-pressure freezing and freeze-substitution. Bacterial biofilms on micro-dialysis tubing or Isopore membrane filters were briefly immersed in 10% glyc-erol or 20% bovine serum albumin in CYE medium. Filters or tubing wastransferred into 200-�m-deep type A aluminum planchettes that were sand-wiched against the flat sides of type B planchettes. The specimens were cryofixedin a Bal-Tec HPM010 high-pressure freezer (2,100 bars, 5 to 7 milliseconds)(Bal-Tec, Inc., Carlsbad, CA). Using the Leica automated freeze-substitutionsystem AFS (Leica Microsystems, Vienna, Austria), cryofixed specimens werefreeze-substituted in anhydrous acetone containing 1% osmium tetroxide and0.1% uranyl acetate and infiltrated with Epon-Araldite following establishedprotocols (34). Specimens were flat-embedded between two microscopy slidesand polymerized at 60°C over 1 to 2 days (35). Resin-embedded biofilm sampleswere remounted under a dissecting microscope for precise orientation.

2D projection transmission electron microscopy for sample surveying. Thin(70- to 100-nm) sections were collected on Formvar-coated slot grids and post-stained with 2% uranyl acetate in 70% methanol followed by either Reynold’s orSato’s lead citrate. The sections were imaged in an FEI Tecnai 12 transmissionelectron microscope (FEI, Eindhoven, The Netherlands) or a Zeiss 10 instru-ment, operated at 120 kV or 100 kV, respectively. These samples were used forultrastructural evaluation as well as to assess the quality of fixation for thetomography studies described below.

Electron microscopy tomography. For tomography, thin sections (�100 nm)were imaged in a Tecnai T20 LaB6 instrument operated at 200 kV (UCSF), andsemithick (200- to 300-nm) sections were imaged in an FEI Tecnai F30 micro-scope operating at 300 kV (Boulder Laboratory for 3D Electron Microscopy ofCells, University of Colorado). Binned 2k by 2k tilt series were collected on 4kby 4k charge-coupled-device Gatan camera (Gatan, Inc., Pleasanton, CA) every1° from �70° to �70°, using the UCSF tomography or SerialEM softwarepackage (32). The nominal setting of defocus was �0.2 �m to �1 �m, and thepixel size of the data corresponded to 1 nm. Series were aligned with the help ofeither 10-nm or 15-nm gold fiducial markers (BBI Research, Inc., WI), and 3Dreconstructed using the IMOD software package (24). IMOD was also used fordata inspection and analysis. Where appropriate, 3D volume data were inspectedinteractively using the volume and surface rendering tools VOLUME ROVER(1) and CHIMERA (39; http://www.cgl.ucsf.edu/chimera).

RESULTS AND DISCUSSION

Structural organization of Myxococcus xanthus biofilms. Tofacilitate specimen handling for freezing and ultrastructuralanalysis, Myxococcus xanthus biofilms were grown on two typesof surfaces: Isopore membranes and cellulose tubes. In bothcases, the bacteria assumed a preferred orientation along thesurface, allowing us to select for either longitudinal (Fig. 1A)or cross-section (Fig. 1B and C) views. The preferred align-ment of the bacterial cells along the surface that they adhered

to may be explained by elasticotaxis (44). The biofilms growingon Isopore filters were noticeably thicker, typically 4 to 10 celllayers thick, whereas the biofilms grown on cellulose tubeswere thin, sometimes as thin as 2 cell layers, and more denselypacked. Biofilms on both supports were exquisitely preservedby the high-pressure freezing and freeze-substitution proce-dures.

We imaged the same areas of M. xanthus biofilms by 2Dprojection and by 3D tomography, and the side-by-side com-parison seen in Fig. 2 illustrates the radical improvement ofresolution in the z axis that is provided by 3D tomography. Thebiofilms grown on Isopore filters were sectioned longitudinally(Fig. 2A and B), and thus the structures of both the cytoplasmand the chromatin body are well defined. In this case, thebacteria are more sparsely populated than on the cellulosetubes, and therefore vesicles in the extracellular space wereeasily visualized. The densely packed biofilm grown on cellu-lose was visualized in cross-section (Fig. 2C and D); cytoplas-mic elements were well resolved, and cell-cell junctions be-tween closely apposed cells were observed.

Organization of bacterial chromosomes. At the center of thebacteria we detected high-contrast multistranded filamentousdensities resembling a twisted closed circle that often showedprominent clefts along the longitudinal axis (Fig. 3A to C). Weinterpret the entity of these filamentous densities as the bac-terial chromosomal DNA. Interestingly, and particularly obvi-ous when viewed in cross-section, we typically found the chro-mosomal DNA at the center of the cell, independent ofwhether or not the cell is in intimate lateral contact with itsneighboring cell. Therefore, even in such close contacts (seebelow), there is no structural evidence of chromosomal DNAexchange, although the exchange of small portions cannot beexcluded. In the reconstructed volumes, strands that we con-sider to be DNA were �3 to 5 nm thick.

Intracellular structures in M. xanthus biofilms. Granularand smooth cytoplasmic inclusions were observed in cross-sections (Fig. 1B and C and 2C and D) and presumably rep-resent poly-�-hydroxybutyric acid storage, although otherforms of energy or lipid storage cannot be ruled out. Interest-

FIG. 1. Ultrastructural characterization of high-pressure-frozen,freeze-substituted Myxococcus xanthus biofilms. (A) 2D projection im-ages of a longitudinal section of thick and sparsely populated biofilmgrown on an Isopore filter, showing intracellular features and vesicles,some of which appear to enclose cargo. (B and C) Cross-sectionsthrough a thin, tightly packed biofilm resting on a cellulose substrate.Notice how the bacteria accommodate each other by adjusting theirshapes. Excellent preservation is manifest in the absence of extractionartifacts in both cellular contents and extracellular material. The bio-film boundary at the air-extracellular material interface is denoted bymultiple arrows. Bar, 500 nm.

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ingly, in addition to these putative poly-�-hydroxybutyricstorage granules, we observed two types of intracellular or-ganelle-like features. While detectable in 2D projections, thecharacteristic shapes of those intracellular structures becameobvious only by 3D tomographic reconstruction. These mem-brane-delineated features resemble organelles and are eitherof crescent (Fig. 3D) or concentric spherical (Fig. 3E) shape.The intermembrane space in the crescent-shaped compart-ment, as well as the sphere-like organelle, is bridged by fila-ments that, given their staining properties, are likely to beproteinaceous in nature. Interestingly, ribosomes or similarlysized macromolecules are excluded from the cytoplasmic spaceenclosed by the rings. The relationship between the sphere-likeand the crescent-like features remains elusive, but thethreaded intermembrane space is a common feature.

Cell-substrate and cell-cell cohesion. In densely populatedthin-layer biofilms, cells were in intimate contact with oneanother. Large areas of cell-cell contacts were already visible in2D projections (Fig. 1B and C and 2C) but were studied ingreater detail by electron tomography (Fig. 2D). 2D projectionviews suggest a zipper-like structure that connects the outermembranes at short segments or along almost the entire cell-cell contact surface (Fig. 4A and B). Between adjacent bacte-rial outer membranes we noticed a small but noticeable gapthroughout the adhesion zone bridged by macromolecules witha size and stain distribution suggestive of a proteinaceous na-ture, indicating that this direct cell-cell interaction is mediated

by outer membrane protein contacts. We often found distinctdeformations of the cell shape to allow for extensive cell-cellcontact (Fig. 1B and C), suggesting that these contacts markdistinct cell-cell interactions and are not simply a result of closepacking of bacteria (for a review of bacterial shapes, see ref-erence 50). In the tightly packed biofilms, EPS and fimbriae donot appear to be involved in these cell-cell contacts. Instead,these cell-cell contacts show ultrastructural similarities to thepreviously described “conjugative” (40) and “conjugational”(10) junctions described for Escherichia coli. Intimate bacterialinteractions have also been seen in M. xanthus cell pairs andmonolayers studied by atomic force microscopy, althoughthese studies did not allow the characterization of such inter-actions (38). Conceivably, these cell-cell junctions play a role inthe swarming phenomenon central to its predatory character-istics. It is noteworthy that despite the fact that the outermembranes are in close proximity, we have not observed directouter membrane fusion, which was postulated by Nudleman etal. to account for the direct cell-to-cell transfer of outer mem-brane proteins (36). While we cannot exclude that such fusionevents can occur, we propose that other means of outer mem-brane protein transfer may exist, e.g., via vesicles, as will bediscussed in more detail below.

Sharp biofilm boundary. 2D projection images of the cross-section views of biofilms on cellulose tubes revealed a sharpdiscontinuity at the edge of the biofilm. When visualized byelectron microscopic tomography, this biofilm border is a con-tinuous high-contrast, sheet-like feature that appears to de-

FIG. 2. 3D electron tomographic reconstruction of Myxococcusxanthus biofilms. (A and B) A 250-nm thick section of multicell layeredbiofilm grown on an Isopore filter. (C and D) A 100-nm thin section of4- to 10-cell layered biofilm grown on cellulose. Shown here are 2Dprojection views (A and C) and corresponding slices through tomo-grams (B and D) at �1-nm thickness. Cellular features obscured bysuperimposition in two dimensions are resolved in the third dimensionin the tomogram, allowing 3D visualization and feature extraction.Asterisks denote chromosomal DNA, and arrows point out the biofilmboundary. Bar, 500 nm.

FIG. 3. 3D reconstruction of intracellular features. (A to C) Chro-mosomal DNA is of high contrast and easily detectable. It was typicallyextended along the cellular axis and coiled along the central axis.(D) Crescent-shaped membranous structure. (E) Concentric-ring-shaped organelle. In both panels D and E, the intermembrane space isthreaded by links of unknown nature. Bar, 100 nm.

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marcate the entire biofilm-air interface (Fig. 1B and C and4C). Virtually every available space within this biofilm borderis occupied by vesicles that are 30 to 60 nm in diameter (Fig. 1Band C), while the space outside the border is devoid of thesevesicles. The boundary structure is a thin (�5-nm) sheet ofosmophilic material (Fig. 4C) that resembles in dimension andappearance a lipid bilayer. The affinity for osmium and itsresulting electron density make it unlikely that this boundarylayer consists of secreted carbohydrates. While we cannot ex-clude that this boundary represents aggregation of denaturedextracellular proteins, it is uniform in thickness and shows astriking resemblance to the lipid bilayer of the bacterial mem-branes in this preparation. Given the number of secreted ves-icles (discussed below) and the fact that lipids naturally accu-mulate at an air-water interface (13, 14, 47), we propose thatthe sharp biofilm boundary consists of a lipid layer that pro-tects the biofilm from desiccation.

Visualization of extracellular filaments: T4P. Extracellularfilamentous materials were not easily detectable in 2D projec-tions, but a set of smoothly curved filaments, �4 to 6 nm indiameter and micrometers in length, became obvious in 3Dreconstructions (Fig. 4D). We interpret these filaments to beT4P, because their dimensions and the structure of their areaof origin in the cell are consistent with those seen in otherultrastructural and atomic force microscopy studies (38).These filaments protrude from a ribosome-devoid polar regionof M. xanthus (Fig. 4D). In serial tomograms we could trace a

set of T4P filaments more than 1 �m into the extracellularspace. TFP are known to mediate cell-substrate interactionsand are crucial for S motility in M. xanthus and other bacteriathat display gliding motility (18). The study presented here didnot allow us to assess the motility of individual bacteria in thebiofilm, but it is interesting to notice that the ratio of filamentbundles, which are found to extend for more than 1 �mthroughout the biofilm, is low compared to the number of cellspresent in such volumes, suggesting that only a few cells as-semble T4P. In conclusion, the role of T4P in biofilms remainsunclear. If the T4P bundle is used for S motility, one wouldexpect the filament to end on a defined substrate or anothernearby cell, several of which were well within reach. Also, if itsrole is to propel the cell forward, one would expect the T4P tobe straightened under tension. Instead, the filaments we ob-served in biofilms are smoothly curved and are even found toassume a direction perpendicular to the original axis. Thescarcity of assembled T4P in the biofilm and the morphology ofvisualized T4P suggest that the role of T4P in a biofilm may bedifferent from their well-documented role in S motility.

Possible roles of extracellular vesicles. The extracellular ma-terial in biofilms is traditionally thought of as being filled withEPS, which serves as an enveloping medium. EPS has beenfound to be essential in M. xanthus communities (27, 28).However, EPS is a term that is loosely used to describe allextracellular material, independent of its actual chemical com-position. In a proteomic study of M. xanthus biofilms, theextracellular material was found to be enriched in proteins ofnovel function, but putative functions could be assigned foronly 5 of the 21 proteins identified (9). Apart from playing astructural role, the extracellular matrix is also likely to mediateinterbacterial signaling and communication. Interestingly, quo-rum signaling involving 2-heptyl-3-hydroxy-4-quinolone waslinked to vesicle formation in P. aeruginosa (29). Extracellularmembrane vesicles have been observed in several planktonicgram-negative bacteria and their biofilm communities (2, 4,33).

In both thin and thick Myxococcus biofilms, the extracellularspace is filled with vesicles with a diameter of 30 to 60 nm (Fig.5). In the thin biofilms, almost every conceivable space is filledwith vesicles, including the space underneath the sharp biofilmboundary (Fig. 1B and C). It should be emphasized that in ourcryo-immobilized samples we did not observe signs of cellularstress and cell disintegration. The membranes are smooth,unlike what is often found in chemically fixed samples, wheremembranes display blebbing and other artifacts. Where bacte-ria were spread at lower density, such as in the thick biofilms,we typically found the vesicles in direct vicinity of bacterialmembranes. 3D reconstruction revealed that vesicles were ei-ther directly in contact with bacterial membranes or linked tocell membranes by what appear to be proteinaceous tethers(Fig. 5B, D, and E). Occasionally, a vesicle was seen in directcontact with three bacterial cell membranes (Fig. 5C). Vesicleswere found tethered not only to the bacterial outer membranesurface but also to each other (Fig. 5D and E). The tethersappear as short filaments of about 3 nm in diameter and tensof nanometers in length. On the surface of a number of vesicleswe find protrusions that are likely to be integral membraneproteins. The majority of vesicles display an internal stain

FIG. 4. 3D reconstruction of extracellular elements in biofilm or-ganization. (A and B) In the thin and highly packed biofilms (Fig. 1Band C), cell-cell contacts are characterized by the membranes in closecontact stitched to one another by extracellular linkages, in a bondstrong enough to overcome the electrostatic repulsion of like surfaces.Interbacterial outer membrane fusion events were not observed.(C) At the air-extracellular material interface, there was a highly os-micated, single-track film along the entire surface of the biofilm (de-noted by arrows), presumably protecting it from desiccation. (D) Thepolar T4P (asterisks) were easily visualized in tomograms, whereasthey were not obvious from projection images. Bar, 100 nm.

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distribution that suggests either a protein cargo or some kindof internal organization.

Membrane vesicles have been shown before by electron mi-croscopy studies (2, 4, 12, 23), where they were pinched offfrom the surface and released into the surrounding medium.However, in some of these studies, membrane blebbing in-duced by chemical fixation artifacts could not be ruled out. Ourresults cannot be attributed to fixation artifacts, since high-pressure freezing occurs instantaneously (within milliseconds).Moreover, membrane vesicles were observed in high-pressure-frozen P. aeruginosa biofilms (42), although at a much lowerdensity, and the intervesicular tethers were not described inthat study. While a variety of possible functions for these ex-tracellular vesicles have been proposed (29, 31, 42), the exactrole of such vesicles remains elusive. We propose that thesevesicles serve as a vehicle for the interbacterial exchange ofouter membrane proteins. Exchange of bacterial outer mem-brane proteins in the absence of gene transfer was shown byNudleman et al. (36), who attributed the exchange to transientfusion of the outer membranes.

In this study we did not detect outer membrane fusion;instead, we observe a wealth of vesicles that are in intimatecontact with the outer membranes of one or several bacteria,suggesting vesicles as the most likely candidate for intercellularprotein transfer. A vesicle-mediated mechanism of proteintransfer, however, implies that bacteria must have a sophisti-cated mechanism not only for vesicle secretion but also for theuptake of the vesicles. We suggest that the protrusions on thevesicle surface represent receptor molecules that are importantin vesicle uptake.

Clearly, vesicles are abundant and are likely to play a role inM. xanthus biofilm physiology. These observations contributeto the foundation of a new field of study, calling for composi-tional analysis and time-lapse high-resolution microscopy stud-

ies, to solve the mystery of bacterial vesicle function, forma-tion, and fusion (30, 31).

Concluding remarks. Microbial biofilms are ubiquitous in allnutrient-sufficient ecosystems (7), and functional mature com-munities develop by processes similar to those that producemulticellular eukaryotic organisms (37). The well-establishedmethods of chemical fixation and 2D projections of thin sec-tions have produced a rich literature concerning the ultrastruc-ture of individual bacterial cells, but transmission electron mi-croscopy studies of intact biofilm communities are relativelyrare. Advancements in high-pressure freezing and freeze-sub-stitution methods have allowed us to minimize fixation arti-facts, or to avoid them altogether, and developments in tomog-raphy allow improved resolution in the z axis. In this study wehave combined these techniques to examine, for the first time,two different types of biofilm communities produced by thesame organism (M. xanthus) on two different surfaces. In thisexploratory study we have visualized cellular structures (e.g.,chromatin and cytoplasmic organelles), and we have resolvedother structures that may enable interactions between the cellsthat comprise biofilm communities. These include cell-cell“zippers,” pili, and vesicles that appear to be tethered to eachother and to cells and to contain resolvable “cargo.”

ACKNOWLEDGMENTS

We thank Reena Zalpuri of the UC Berkeley Electron MicroscopyLaboratory; the Boulder Laboratory for 3D Electron Microscopy ofCells (BL3DEMC), University of Colorado; and Michael Braunfeldand David Agard at UCSF.

This project was supported by the Director, Office of Science, of theU.S. Department of Energy under contract DE-AC03-76SF00098 toM. Auer; by National Institutes of Health grant GM54666 to W. Shi;and by an Amado Foundation grant to J. W. Costerton. TheBL3DEMC is supported by a grant from the National Center forResearch Resources of the National Institutes of Health (RR-00592)to A. Hoenger.

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FIG. 5. The biofilm extracellular space is filled with vesicles. Thevesicles are tethered to the bacterial membrane, and to each other, andare arranged linearly in a nonrandom fashion. (A) The indentation inthe bacterial membrane may suggest either an endo- or exocytosisevent. (A, B, and D) The vesicle interior displays densities that areindicative of either the presence of a cargo or an underlying proteinorganization. The vesicle is tethered to the membrane by at least oneor two links of unknown nature. (C) Occasionally, vesicles were foundin simultaneous contact with three bacterial outer membranes.(E) Five vesicles shown in linear arrangement close to the bacterial cellsurface. Links between the vesicles and between vesicles and the bac-terial surface were observed. Bar, 100 nm.

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