Three-Dimensional Macromolecular Organization of Cryofixed Myxococcus xanthus Biofilms as Revealed by Electron Microscopic Tomography

Abstract

Despite the fact that most bacteria grow in biofilms in natural and pathogenic ecosystems, very little is known 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, sample aggregation, and sample extraction. As a further innovation we have, for the first time in biofilm research, used electron tomography and three-dimensional (3D) visualization to better resolve the macromolecular 3D ultrastructure of a biofilm. This combination of superb specimen preparation and greatly improved resolution in the z axis has opened a window in studies of Myxococcus xanthus cell ultrastructure and biofilm community architecture. New structural information on the chromatin body, cytoplasmic organization, membrane apposition between adjacent cells, and structure and distribution of pili and vesicles in the biofilm matrix is presented.

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

While a variety of biofilms have been studied in depth by optical imaging approaches, only a few have been faithfully preserved and visualized at high resolution by transmission electron microscopy (3, 22, 25, 26, 45, 48, 49). A recent ultrastructural study of Pseudomonas aeruginosa biofilms shows that high-pressure freezing/freeze-substitution results in superior preservation (22). Unlike conventional sample preparation, millisecond cryofixation and low-temperature dehydration protocols minimize artifacts from chemical fixation, extraction, and aggregation (15, 16, 19, 20, 22).

Conventional two-dimensional (2D) imaging provides x,y positional information but cannot resolve features along the z direction. Conventional 2D projection imaging of thin (∼70- to 100-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 acquisition and 3D reconstruction, upon which both intracellular and extracellular features are resolved. In this way, cellular organelles, molecular machines, and macromolecular complexes can be visualized in their native microenvironment. To date, electron tomographic 3D reconstruction of a bacterial biofilm has not been reported.

Myxococcus xanthus is a well-studied gram-negative soil scavenger and predator (21), and it serves as a model system for biofilm formation. M. xanthus moves over surfaces either by adventurous (A) motility or social (S) motility (for a review, see reference 18). It forms spore-bearing fruiting bodies upon starvation, in a process that requires the coordinated movement of many thousands of cells. S motility relies on retraction of type IV pili (T4P) upon interaction with amine-containing polysaccharides in the extracellular matrix (11, 27, 38, 46). T4P are also required for cell surface adhesion and “clumping.” It is unclear how prominent T4P are in stable microbial communities and what their possible role might be. Also, it is unknown how bacterial community members interact with one another and what mechanisms they employ for effective communication and signal and/or material transfer in such dense communities.

Here, we report faithful preservation of M. xanthus biofilms and the first macromolecular resolution-based insights into their 3D architecture, obtained by electron tomographic imaging of rapidly frozen, freeze-substituted, and resin-embedded biofilm sections. The tomograms reveal the 3D organization of the bacterial chromosome, direct cell-cell and cell-substrate interactions, the presence of extracellular filaments (including but not restricted to T4P), and abundant extracellular vesicles that, interestingly, we found to be tethered to each other and to the bacterial outer membrane.

MATERIALS AND METHODS

Ten milliliters of CYE broth, consisting of 5 mM MOPS (morpholinepropanesulfonic 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 this culture was used to inoculate 200 ml of CYE broth, which was incubated for 18 h at 32°C at 100 rpm. The culture was concentrated to a final optical density at 600 nm 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 of Isopore membrane filters (HTTP, 0.4 μm; Millipore, MA), and subsequently placed on CYE agar. Ten microliters of concentrated culture (optical density at 600 nm of 10) was placed on top of each Spectra/Por fiber/Isopore filter or Isopore membranes and incubated for 6 h at 32°C followed by 20 h at room temperature. In the case of cellulose tubes, biofilms were grown by placing the ∼200-μm-thick cellulose microdialysis tube with the Myxococcus culture on top of a moist agar plate prior to overnight shipment. Using these culturing conditions, the biofilms growing on Isopore filters were noticeably thicker, typically 4 to 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 microdialysis tubing or Isopore membrane filters were briefly immersed in 10% glycerol or 20% bovine serum albumin in CYE medium. Filters or tubing was transferred into 200-μm-deep type A aluminum planchettes that were sandwiched against the flat sides of type B planchettes. The specimens were cryofixed in a Bal-Tec HPM010 high-pressure freezer (2,100 bars, 5 to 7 milliseconds) (Bal-Tec, Inc., Carlsbad, CA). Using the Leica automated freeze-substitution system AFS (Leica Microsystems, Vienna, Austria), cryofixed specimens were freeze-substituted in anhydrous acetone containing 1% osmium tetroxide and 0.1% uranyl acetate and infiltrated with Epon-Araldite following established protocols (34). Specimens were flat-embedded between two microscopy slides and polymerized at 60°C over 1 to 2 days (35). Resin-embedded biofilm samples were 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 poststained with 2% uranyl acetate in 70% methanol followed by either Reynold's or Sato's lead citrate. The sections were imaged in an FEI Tecnai 12 transmission electron microscope (FEI, Eindhoven, The Netherlands) or a Zeiss 10 instrument, operated at 120 kV or 100 kV, respectively. These samples were used for ultrastructural evaluation as well as to assess the quality of fixation for the tomography 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), and semithick (200- to 300-nm) sections were imaged in an FEI Tecnai F30 microscope operating at 300 kV (Boulder Laboratory for 3D Electron Microscopy of Cells, University of Colorado). Binned 2k by 2k tilt series were collected on 4k by 4k charge-coupled-device Gatan camera (Gatan, Inc., Pleasanton, CA) every 1° from −70° to +70°, using the UCSF tomography or SerialEM software package (32). The nominal setting of defocus was −0.2 μm to −1 μm, and the pixel size of the data corresponded to 1 nm. Series were aligned with the help of either 10-nm or 15-nm gold fiducial markers (BBI Research, Inc., WI), and 3D reconstructed using the IMOD software package (24). IMOD was also used for data inspection and analysis. Where appropriate, 3D volume data were inspected interactively 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.

To facilitate specimen handling for freezing and ultrastructural analysis, Myxococcus xanthus biofilms were grown on two types of surfaces: Isopore membranes and cellulose tubes. In both cases, the bacteria assumed a preferred orientation along the surface, allowing us to select for either longitudinal (Fig. 1A) or cross-section (Fig. 1B and C) views. The preferred alignment of the bacterial cells along the surface that they adhered to may be explained by elasticotaxis (44). The biofilms growing on Isopore filters were noticeably thicker, typically 4 to 10 cell layers thick, whereas the biofilms grown on cellulose tubes were thin, sometimes as thin as 2 cell layers, and more densely packed. Biofilms on both supports were exquisitely preserved by the high-pressure freezing and freeze-substitution procedures.

FIG. 1.

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

We imaged the same areas of M. xanthus biofilms by 2D projection and by 3D tomography, and the side-by-side comparison seen in Fig. 2 illustrates the radical improvement of resolution in the z axis that is provided by 3D tomography. The biofilms grown on Isopore filters were sectioned longitudinally (Fig. 2A and B), and thus the structures of both the cytoplasm and the chromatin body are well defined. In this case, the bacteria are more sparsely populated than on the cellulose tubes, and therefore vesicles in the extracellular space were easily visualized. The densely packed biofilm grown on cellulose was visualized in cross-section (Fig. 2C and D); cytoplasmic elements were well resolved, and cell-cell junctions between closely apposed cells were observed.

FIG. 2.

3D electron tomographic reconstruction of Myxococcus xanthus biofilms. (A and B) A 250-nm thick section of multicell layered biofilm grown on an Isopore filter. (C and D) A 100-nm thin section of 4- to 10-cell layered biofilm grown on cellulose. Shown here are 2D projection views (A and C) and corresponding slices through tomograms (B and D) at ∼1-nm thickness. Cellular features obscured by superimposition in two dimensions are resolved in the third dimension in the tomogram, allowing 3D visualization and feature extraction. Asterisks denote chromosomal DNA, and arrows point out the biofilm boundary. Bar, 500 nm.

Organization of bacterial chromosomes.

At the center of the bacteria we detected high-contrast multistranded filamentous densities resembling a twisted closed circle that often showed prominent clefts along the longitudinal axis (Fig. 3A to C). We interpret the entity of these filamentous densities as the bacterial chromosomal DNA. Interestingly, and particularly obvious when viewed in cross-section, we typically found the chromosomal DNA at the center of the cell, independent of whether or not the cell is in intimate lateral contact with its neighboring cell. Therefore, even in such close contacts (see below), there is no structural evidence of chromosomal DNA exchange, although the exchange of small portions cannot be excluded. In the reconstructed volumes, strands that we consider to be DNA were ∼3 to 5 nm thick.

FIG. 3.

3D reconstruction of intracellular features. (A to C) Chromosomal DNA is of high contrast and easily detectable. It was typically extended 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 is threaded by links of unknown nature. Bar, 100 nm.

Intracellular structures in M. xanthus biofilms.

Granular and smooth cytoplasmic inclusions were observed in cross-sections (Fig. 1B and C and 2C and D) and presumably represent poly-β-hydroxybutyric acid storage, although other forms of energy or lipid storage cannot be ruled out. Interestingly, in addition to these putative poly-β-hydroxybutyric storage granules, we observed two types of intracellular organelle-like features. While detectable in 2D projections, the characteristic shapes of those intracellular structures became obvious only by 3D tomographic reconstruction. These membrane-delineated features resemble organelles and are either of crescent (Fig. 3D) or concentric spherical (Fig. 3E) shape. The intermembrane space in the crescent-shaped compartment, as well as the sphere-like organelle, is bridged by filaments that, given their staining properties, are likely to be proteinaceous in nature. Interestingly, ribosomes or similarly sized macromolecules are excluded from the cytoplasmic space enclosed by the rings. The relationship between the sphere-like and the crescent-like features remains elusive, but the threaded intermembrane space is a common feature.

Cell-substrate and cell-cell cohesion.

In densely populated thin-layer biofilms, cells were in intimate contact with one another. Large areas of cell-cell contacts were already visible in 2D projections (Fig. 1B and C and 2C) but were studied in greater detail by electron tomography (Fig. 2D). 2D projection views suggest a zipper-like structure that connects the outer membranes at short segments or along almost the entire cell-cell contact surface (Fig. 4A and B). Between adjacent bacterial outer membranes we noticed a small but noticeable gap throughout the adhesion zone bridged by macromolecules with a size and stain distribution suggestive of a proteinaceous nature, indicating that this direct cell-cell interaction is mediated by outer membrane protein contacts. We often found distinct deformations of the cell shape to allow for extensive cell-cell contact (Fig. 1B and C), suggesting that these contacts mark distinct cell-cell interactions and are not simply a result of close packing of bacteria (for a review of bacterial shapes, see reference 50). In the tightly packed biofilms, EPS and fimbriae do not appear to be involved in these cell-cell contacts. Instead, these cell-cell contacts show ultrastructural similarities to the previously described “conjugative” (40) and “conjugational” (10) junctions described for Escherichia coli. Intimate bacterial interactions have also been seen in M. xanthus cell pairs and monolayers studied by atomic force microscopy, although these studies did not allow the characterization of such interactions (38). Conceivably, these cell-cell junctions play a role in the swarming phenomenon central to its predatory characteristics. It is noteworthy that despite the fact that the outer membranes are in close proximity, we have not observed direct outer membrane fusion, which was postulated by Nudleman et al. to account for the direct cell-to-cell transfer of outer membrane proteins (36). While we cannot exclude that such fusion events can occur, we propose that other means of outer membrane protein transfer may exist, e.g., via vesicles, as will be discussed in more detail below.

FIG. 4.

3D reconstruction of extracellular elements in biofilm organization. (A and B) In the thin and highly packed biofilms (Fig. 1B and C), cell-cell contacts are characterized by the membranes in close contact stitched to one another by extracellular linkages, in a bond strong 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 osmicated, single-track film along the entire surface of the biofilm (denoted by arrows), presumably protecting it from desiccation. (D) The polar T4P (asterisks) were easily visualized in tomograms, whereas they were not obvious from projection images. Bar, 100 nm.

Sharp biofilm boundary.

2D projection images of the cross-section views of biofilms on cellulose tubes revealed a sharp discontinuity at the edge of the biofilm. When visualized by electron microscopic tomography, this biofilm border is a continuous high-contrast, sheet-like feature that appears to demarcate the entire biofilm-air interface (Fig. 1B and C and 4C). Virtually every available space within this biofilm border is occupied by vesicles that are 30 to 60 nm in diameter (Fig. 1B and C), while the space outside the border is devoid of these vesicles. The boundary structure is a thin (∼5-nm) sheet of osmophilic material (Fig. 4C) that resembles in dimension and appearance a lipid bilayer. The affinity for osmium and its resulting electron density make it unlikely that this boundary layer consists of secreted carbohydrates. While we cannot exclude that this boundary represents aggregation of denatured extracellular proteins, it is uniform in thickness and shows a striking resemblance to the lipid bilayer of the bacterial membranes in this preparation. Given the number of secreted vesicles (discussed below) and the fact that lipids naturally accumulate at an air-water interface (13, 14, 47), we propose that the sharp biofilm boundary consists of a lipid layer that protects the biofilm from desiccation.

Visualization of extracellular filaments: T4P.

Extracellular filamentous materials were not easily detectable in 2D projections, but a set of smoothly curved filaments, ∼4 to 6 nm in diameter and micrometers in length, became obvious in 3D reconstructions (Fig. 4D). We interpret these filaments to be T4P, because their dimensions and the structure of their area of origin in the cell are consistent with those seen in other ultrastructural and atomic force microscopy studies (38). These filaments protrude from a ribosome-devoid polar region of M. xanthus (Fig. 4D). In serial tomograms we could trace a set of T4P filaments more than 1 μm into the extracellular space. TFP are known to mediate cell-substrate interactions and are crucial for S motility in M. xanthus and other bacteria that display gliding motility (18). The study presented here did not allow us to assess the motility of individual bacteria in the biofilm, but it is interesting to notice that the ratio of filament bundles, which are found to extend for more than 1 μm throughout the biofilm, is low compared to the number of cells present in such volumes, suggesting that only a few cells assemble T4P. In conclusion, the role of T4P in biofilms remains unclear. If the T4P bundle is used for S motility, one would expect the filament to end on a defined substrate or another nearby cell, several of which were well within reach. Also, if its role is to propel the cell forward, one would expect the T4P to be straightened under tension. Instead, the filaments we observed in biofilms are smoothly curved and are even found to assume a direction perpendicular to the original axis. The scarcity of assembled T4P in the biofilm and the morphology of visualized T4P suggest that the role of T4P in a biofilm may be different from their well-documented role in S motility.

Possible roles of extracellular vesicles.

The extracellular material in biofilms is traditionally thought of as being filled with EPS, which serves as an enveloping medium. EPS has been found to be essential in M. xanthus communities (27, 28). However, EPS is a term that is loosely used to describe all extracellular material, independent of its actual chemical composition. In a proteomic study of M. xanthus biofilms, the extracellular material was found to be enriched in proteins of novel function, but putative functions could be assigned for only 5 of the 21 proteins identified (9). Apart from playing a structural role, the extracellular matrix is also likely to mediate interbacterial signaling and communication. Interestingly, quorum signaling involving 2-heptyl-3-hydroxy-4-quinolone was linked to vesicle formation in P. aeruginosa (29). Extracellular membrane vesicles have been observed in several planktonic gram-negative bacteria and their biofilm communities (2, 4, 33).

In both thin and thick Myxococcus biofilms, the extracellular space is filled with vesicles with a diameter of 30 to 60 nm (Fig. 5). In the thin biofilms, almost every conceivable space is filled with vesicles, including the space underneath the sharp biofilm boundary (Fig. 1B and C). It should be emphasized that in our cryo-immobilized samples we did not observe signs of cellular stress and cell disintegration. The membranes are smooth, unlike what is often found in chemically fixed samples, where membranes display blebbing and other artifacts. Where bacteria were spread at lower density, such as in the thick biofilms, we typically found the vesicles in direct vicinity of bacterial membranes. 3D reconstruction revealed that vesicles were either directly in contact with bacterial membranes or linked to cell membranes by what appear to be proteinaceous tethers (Fig. 5B, D, and E). Occasionally, a vesicle was seen in direct contact with three bacterial cell membranes (Fig. 5C). Vesicles were found tethered not only to the bacterial outer membrane surface but also to each other (Fig. 5D and E). The tethers appear as short filaments of about 3 nm in diameter and tens of nanometers in length. On the surface of a number of vesicles we find protrusions that are likely to be integral membrane proteins. The majority of vesicles display an internal stain distribution that suggests either a protein cargo or some kind of internal organization.

FIG. 5.

The biofilm extracellular space is filled with vesicles. The vesicles are tethered to the bacterial membrane, and to each other, and are arranged linearly in a nonrandom fashion. (A) The indentation in the bacterial membrane may suggest either an endo- or exocytosis event. (A, B, and D) The vesicle interior displays densities that are indicative of either the presence of a cargo or an underlying protein organization. The vesicle is tethered to the membrane by at least one or two links of unknown nature. (C) Occasionally, vesicles were found in simultaneous contact with three bacterial outer membranes. (E) Five vesicles shown in linear arrangement close to the bacterial cell surface. Links between the vesicles and between vesicles and the bacterial surface were observed. Bar, 100 nm.

Membrane vesicles have been shown before by electron microscopy studies (2, 4, 12, 23), where they were pinched off from the surface and released into the surrounding medium. However, in some of these studies, membrane blebbing induced by chemical fixation artifacts could not be ruled out. Our results 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 lower density, and the intervesicular tethers were not described in that study. While a variety of possible functions for these extracellular vesicles have been proposed (29, 31, 42), the exact role of such vesicles remains elusive. We propose that these vesicles serve as a vehicle for the interbacterial exchange of outer membrane proteins. Exchange of bacterial outer membrane proteins in the absence of gene transfer was shown by Nudleman et al. (36), who attributed the exchange to transient fusion of the outer membranes.

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

Clearly, vesicles are abundant and are likely to play a role in M. xanthus biofilm physiology. These observations contribute to the foundation of a new field of study, calling for compositional analysis and time-lapse high-resolution microscopy studies, to solve the mystery of bacterial vesicle function, formation, and fusion (30, 31).

Concluding remarks.

Microbial biofilms are ubiquitous in all nutrient-sufficient ecosystems (7), and functional mature communities develop by processes similar to those that produce multicellular eukaryotic organisms (37). The well-established methods of chemical fixation and 2D projections of thin sections have produced a rich literature concerning the ultrastructure of individual bacterial cells, but transmission electron microscopy studies of intact biofilm communities are relatively rare. Advancements in high-pressure freezing and freeze-substitution methods have allowed us to minimize fixation artifacts, or to avoid them altogether, and developments in tomography allow improved resolution in the z axis. In this study we have combined these techniques to examine, for the first time, two different types of biofilm communities produced by the same organism (M. xanthus) on two different surfaces. In this exploratory study we have visualized cellular structures (e.g., chromatin and cytoplasmic organelles), and we have resolved other structures that may enable interactions between the cells that comprise biofilm communities. These include cell-cell “zippers,” pili, and vesicles that appear to be tethered to each other and to cells and to contain resolvable “cargo.”