Communication among oral bacteria

Abstract

Human oral bacteria interact with their environment by attaching to surfaces and establishing mixed-species communities. As each bacterial cell attaches, it forms a new surface to which other cells can adhere. Adherence and community development are spatiotemporal; such order requires communication. The discovery of soluble signals, such as autoinducer-2, that may be exchanged within multispecies communities to convey information between organisms has emerged as a new research direction. Direct-contact signals, such as adhesins and receptors, that elicit changes in gene expression after cell-cell contact and biofilm growth are also an active research area. Considering that the majority of oral bacteria are organized in dense three-dimensional biofilms on teeth, confocal microscopy and fluorescently labeled probes provide valuable approaches for investigating the architecture of these organized communities in situ. Oral biofilms are readily accessible to microbiologists and are excellent model systems for studies of microbial communication. One attractive model system is a saliva-coated flowcell with oral bacterial biofilms growing on saliva as the sole nutrient source; an intergeneric mutualism is discussed. Several oral bacterial species are amenable to genetic manipulation for molecular characterization of communication both among bacteria and between bacteria and the host. A successful search for genes critical for mixed-species community organization will be accomplished only when it is conducted with mixed-species communities.

Figures

FIG. 1.

Spatiotemporal model of oral bacterial colonization, showing recognition of salivary pellicle receptors by early colonizing bacteria and coaggregations between early colonizers, fusobacteria, and late colonizers of the tooth surface. Each coaggregation depicted is known to occur in a pairwise test. Collectively, these interactions are proposed to represent development of dental plaque and are redrawn from Kolenbrander and London (79). Starting at the bottom, primary colonizers bind via adhesins (round-tipped black line symbols) to complementary salivary receptors (blue-green vertical round-topped columns) in the acquired pellicle coating the tooth surface. Secondary colonizers bind to previously bound bacteria. Sequential binding results in the appearance of nascent surfaces that bridge with the next coaggregating partner cell. Several kinds of coaggregations are shown as complementary sets of symbols of different shapes. One set is depicted in the box at the top. Proposed adhesins (symbols with a stem) represent cell surface components that are heat inactivated (cell suspension heated to 85°C for 30 min) and protease sensitive; their complementary receptors (symbols without a stem) are unaffected by heat or protease. Identical symbols represent components that are functionally similar but may not be structurally identical. Rectangular symbols represent lactose-inhibitable coaggregations. Other symbols represent components that have no known inhibitor. The bacterial strains shown are Actinobacillus actinomycetemcomitans, Actinomyces israelii, Actinomyces naeslundii, Capnocytophaga gingivalis, Capnocytophaga ochracea, Capnocytophaga sputigena, Eikenella corrodens, Eubacterium spp., Fusobacterium nucleatum, Haemophilus parainfluenzae, Porphyromonas gingivalis, Prevotella denticola, Prevotella intermedia, Prevotella loescheii, Propionibacterium acnes, Selenomonas flueggei, Streptococcus gordonii, Streptococcus mitis, Streptococcus oralis, Streptococcus sanguis, Treponema spp., and Veillonella atypica.

FIG. 2.

Human oral biofilm formed in vitro with a saliva inoculum and using sterile saliva as its sole source of nutrient. The 25-μm-thick biofilm was grown overnight suspended from the underside of the coverslip of a flowcell with saliva flowing through once at 0.2 ml per min. Bacterial juxtaposition and biofilm architecture were imaged by confocal scanning laser microscopy after staining the cells with Live/Dead stain (Molecular Probes, Eugene, Oreg.). The color of the cells is from the red (propidium iodide; damaged or permeable cell membrane) and green (SYTO 9; healthy cell) fluorescent stains. Colocalization of both fluorophores results in yellow staining. Confocal scanning laser microscopy acquires optical sections through the biofilm; each optical section is 0.5 μm thick. The entire biofilm is represented in six images (A to F). Panel A is the 0.5-μm optical section at the substratum and shows the biofilm footprint. Panel F is the top 0.5 μm of the biofilm where it projects into the lumen of the flowcell. The other four projection images contain eight sections per projection and show the 4-μm-thick regions from 4 to 8 μm from the substratum (B), 8 to 12 μm from the substratum (C), 12 to 16 μm from the substratum (D), and 16 to 20 μm from the substratum (E). Regions indicated by arrows are described in the text. Bar, 10 μm. Microscopic observations and image acquisition were performed on a TCS 4D system (Leica Lasertechnik GmbH, Heidelberg, Germany).

FIG. 3.

Four-genus mixture of oral bacteria stained with a specific probe by FISH and the nonspecific nucleic acid stain SYTO 59. (A) Confocal micrograph of a field of cells stained with the nucleic acid stain SYTO 59. The field of cells contains the following species: Actinomyces serovar WVA963 strain PK1259 (a), F. nucleatum PK1594 (f), S. gordonii DL1 (s), and Veillonella atypica PK1910 (v). (B) Confocal micrograph of the same field showing the location of the fluorescein isothiocyanate-labeled actinomyces-specific probe. The image demonstrates that the probe interacts only with actinomyces cells (a). (C) Overlay of confocal micrographs (A and B), demonstrating the specificity of the actinomyces probe. Areas of colocalization of fluorescein isothiocyanate and SYTO-59 markers appear yellow. The yellow actinomyces cells are in contact with other cells seen at the edges of the actinomyces cluster. Coaggregations (c1 to c5) of different species within the mixed culture are also visible. (D) Differential interference contrast image of the field of cells using transmitted light. The distinct morphologies of the various cells in the mixed culture are visible. Bar, 10 μm. Microscopic observations and image acquisition were performed on a TCS 4D system (Leica Lasertechnik GmbH, Heidelberg, Germany).

FIG. 4.

Representative confocal scanning laser microscopy images of a two-species biofilm formed by S. oralis 34 and A. naeslundii T14V. Maximum projection images were taken at 0 h (A and B) and 18 h (C and D) of salivary flow. S. oralis 34 was incubated statically in a saliva-coated flowcell for 15 min before initiation of salivary flow at 0.2 ml/min for 15 min. A. naeslundii T14V was then added and incubated statically for 15 min, and salivary flow was resumed for 15 min (equals time zero). S. oralis 34 cells were labeled with rabbit anti-S. oralis 34 serum (gift of J. Cisar), followed by indodicarbocyanine-conjugated goat anti-rabbit immunoglobulin antibody (Jackson ImmunoResearch Laboratories, Inc., West Grove, Pa.); indodicarbocyanine fluorescence is presented in green. A. naeslundii T14V was labeled with a mouse monoclonal antibody against type 1 fimbriae (gift of J. Cisar), followed by indocarbocyanine-conjugated goat anti-mouse immunoglobulin antibody (Jackson ImmunoResearch Laboratories); indocarbocyanine fluorescence is presented in red. A. naeslundii cells are frequently located in direct proximity to S. oralis cells. After 18 h of saliva flow, growth of both genera is apparent. Dimensions of the regions displayed are 250 μm by 250 μm (x-y perspectives; A and C) and 83 μm by 83 μm (x-y perspectives; B and D; 3× zoom of the center portion of the upper panels). Rotation of the maximum projection to display the x-z perspective (83 μm by 10 μm) is shown below panels B and D. The substratum position in the x-z perspective is indicated by the white line. Bars, 20 μm (A and C) and 10 μm (B and D). Microscopic observations and image acquisition were performed on a TCS 4D system (Leica Lasertechnik GmbH, Heidelberg, Germany).

FIG. 5.

Alignment of the deduced amino acid sequences of luxS from S. gordonii DL1 (Sg; accession number AY081773), S. mutans UA159 (Sm; www.genome.ou.edu/smutans.html), A. actinomycetemcomitans HK1651 (Aa; www.genome.ou.edu/act.html), V. harveyi BB120 (Vh; accession no. AF120098), and P. gingivalis W83 (Pg; www.tigr.org). Consensus of at least 50% identical amino acid residues is denoted by black boxes; conserved amino acid substitutions are highlighted with gray boxes. Asterisks are placed above the 23 amino acid residues that were shown to be invariant in 26 LuxS sequences aligned by Hilgers and Ludwig (64).

FIG. 6.

(A) Involvement of Ag I/II family members in Streptococcus-Actinomyces coaggregations. The diagram shows coaggregations between S. gordonii DL1, S. oralis H1, and S. oralis 34 (representatives of streptococcal coaggregation groups 1, 2, and 3, respectively) with six actinomyces coaggregation groups (groups A, B, C, D, E, and F) (72). Actinomyces coaggregation groups are symbolized by oblong cellular shapes containing letters representing the groups and surround the three circles representing streptococcal strains DL1, H1, and 34. The model of S. gordonii DL1 coaggregation is based on identification of the independent coaggregation functions of SspA and SspB of S. gordonii DL1 (41). The models for S. oralis strains H1 and 34 are based on pairwise coaggregations between actinomyces and streptococci, lactose-inhibitable coaggregations, and loss of certain coaggregation functions by spontaneous mutants (72). The complementary symbols shown here are described in the legend to Fig. 1. *, putative protein designations. (B) Graphic representation of sspA and sspB (accession number U40027), with divergent regions shown in green and purple and gene fragments amplified from S. oralis 34 and S. oralis H1 encoding Ag I/II homologues. The locations of primers (solid arrowheads) used to amplify the homologous fragments from S. oralis H1 (purple-stippled rectangle) and S. oralis 34 (green-stippled rectangle) are shown. Blast2 alignments (145), shown below, reveal that the divergent regions of S. gordonii sspB and the S. oralis H1 homologue are conserved over the length of this region (conserved sequences appear as a straight line at a 45o angle). However, alignment of the sspA divergent region and the corresponding region from the S. oralis 34 homologue reveals gaps in the sequence identity. The similarities and differences in these sequences may be responsible for the coaggregations with A. naeslundii strains shown in panel A. See text for more detail.