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Appl Environ Microbiol. 2014 November; 80(21): 6583–6590.
PMCID: PMC4249035

Probing of Microbial Biofilm Communities for Coadhesion Partners

A. M. Spormann, Editor

Abstract

Investigations of interbacterial adhesion in dental plaque development are currently limited by the lack of a convenient assay to screen the multitude of species present in oral biofilms. To overcome this limitation, we developed a solid-phase fluorescence-based screening method to detect and identify coadhesive partner organisms in mixed-species biofilms. The applicability of this method was demonstrated using coaggregating strains of type 2 fimbrial adhesin-bearing actinomyces and receptor polysaccharide (RPS)-bearing streptococci. Specific adhesin/receptor-mediated coadhesion was detected by overlaying bacterial strains immobilized to a nitrocellulose membrane with a suspended, fluorescein-labeled bacterial partner strain. Coadhesion was comparable regardless of which cell type was labeled and which was immobilized. Formaldehyde treatment of bacteria, either in suspension or immobilized on nitrocellulose, abolished actinomyces type 2 fimbrial adhesin but not streptococcal RPS function, thereby providing a simple method for assigning complementary adhesins and glycan receptors to members of a coadhering pair. The method's broader applicability was shown by overlaying colony lifts of dental plaque biofilm cultures with fluorescein-labeled strains of type 2 fimbriated Actinomyces naeslundii or RPS-bearing Streptococcus oralis. Prominent coadhesion partners included not only streptococci and actinomyces, as expected, but also other bacteria not identified in previous coaggregation studies, such as adhesin- or receptor-bearing strains of Neisseria pharyngitis, Rothia dentocariosa, and Kingella oralis. The ability to comprehensively screen complex microbial communities for coadhesion partners of specific microorganisms opens a new approach in studies of dental plaque and other mixed-species biofilms.

INTRODUCTION

Oral microbial biofilms have been a focus of long-term interest because of their involvement in dental, oral, and perhaps systemic disease (1, 2). Interbacterial adhesion is thought to play an important role in the spatial development of these biofilms and formation of microbial communities that efficiently utilize available nutrients or resist environmental challenges (3). Recent studies of the oral microbiome (4, 5) revealed a remarkable diversity of bacterial species inhabiting the oral cavity and also showed associations of certain species with oral health or disease (2). Now that the numerous players have been identified, it needs to become better understood how they act together in a concerted fashion in vivo. However, much of the knowledge available to date about interbacterial adhesion is still based on in vitro studies of isolated strains available from culture collections.

In past studies of interbacterial adhesion, an in-suspension coaggregation assay has commonly been used, in which a pair of bacterial cell types are mixed together in suspension and scored for the occurrence of visible aggregates (6). Through the use of such assays, numerous laboratory strains and isolates of human oral bacteria have been tested for their ability to coaggregate with each other in vitro (7). For some of those coaggregating partner strains, the mechanisms of interbacterial adhesion have been studied down to the molecular level by unraveling the structural details of bacterial adhesins and complementary binding sites (8, 9). Also, the temporal succession of bacterial genera has been demonstrated through ex vivo studies of oral biofilms formed on retrievable enamel chips during incubation in the oral environment (10, 11). Using this experimental model, bacterial adhesins and complementary receptors, such as actinomyces type 2 fimbriae and streptococcal receptor polysaccharides (RPS), were colocalized on closely associated bacteria in microbial consortia formed in vivo (12). However, even these studies are restricted by their very nature to known strains of coadhering bacteria and already characterized adhesin and receptor molecules for which molecular probes exist.

The currently available assays for interbacterial adhesion are not well suited to handle the multitude of microbial species present in oral biofilms, many of which assumedly express unknown bacterial adhesin and receptor molecules. For this reason, we have adapted here the bacterial overlay method (13, 14) and combined it with a method for immobilizing bacteria (15) to allow efficient screening of multispecies biofilms for coadhesion partners with the option to isolate and identify them subsequently. The resulting coadhesion assay was first validated using strains of oral actinomyces and streptococci that are known to coaggregate in vitro (8, 9) and engage in coadhesion in vivo (12). As a proof of principle, the assay was then used to screen mixed dental biofilms for unknown coadhesion partners.

MATERIALS AND METHODS

Bacterial culture.

Bacteria used in this investigation included Actinomyces naeslundii WVU45 and type 2 fimbriae-deficient mutant strain WVU45M (16), Streptococcus oralis 34 and RPS-deficient mutant 34M (17), as well as Streptococcus gordonii 38 and the RPS-deficient mutant strain XC3 (18). These bacteria, as well as others isolated from samples of dental plaque (see below), were grown in brain heart infusion (BHI) broth (Difco) as previously described (19) or on plates of Columbia agar (Merck, Darmstadt, Germany) containing 8% defibrinated sheep blood. All strains were maintained as frozen stocks at −80°C using the Cryobank system (Mast Diagnostica GmbH, Reinfeld, Germany) according to the instructions of the manufacturer.

Collection of dental plaque samples.

This study was approved by the Ethics Committee of the Medical School of the University of Regensburg. Mature dental plaque was collected using a sterile scaler (Taylor scaler; Hu-Friedy Mfg. Co., Inc., Tuttlingen, Germany) from interproximal sites of anterior and posterior teeth within each quadrant of the dentition, avoiding touching the gingival tissue. For the collection of early dental plaque, volunteers were instructed to clean their teeth the night before collection and to abstain thereafter from eating, drinking, and brushing teeth before the collection of plaque the next morning. Early dental plaque was harvested by rubbing the anterior surfaces of the upper and lower incisors with sterile cotton swabs. Bacteria were then suspended by swirling the swabs in 200 μl of sterile saline solution. The resulting suspension was vortex mixed, and serial 10-fold dilutions were plated on media, as described above, followed by incubation at 37°C in 5% CO2 for 1 or 2 days for the growth of isolated colonies.

Coadhesion assay.

Bacteria harvested from overnight BHI broth cultures were washed with phosphate-buffered saline (PBS), adjusted to an optical density at 600 nm (OD600) of 1.0 (~108 CFU/ml), and labeled with fluorescein isothiocyanate as previously described (14). Where indicated, labeled bacteria were incubated in 15% formaldehyde for 1 h to denature bacterial adhesins and then washed four times with 10 ml of PBS. Suspension-grown bacteria to be used for surface immobilization were washed in PBS before transfer to nitrocellulose membranes (Protran BA 85; Schleicher & Schuell, Germany) using a dot-blotting apparatus (Minifold 1; Schleicher & Schuell) connected to a vacuum pump (ME 4R; Vacuubrand). Volumes of 100 μl of bacterial suspensions in serial 2-fold dilutions, starting with a concentration of 108 CFU/ml, were added to the wells. Once the fluid of the suspensions was drawn through, the wells were twice flushed with 100 μl of PBS each. After disassembly of the dot blot apparatus, the nitrocellulose membrane carrying the immobilized bacteria was rinsed once and then washed two times with Tris-HCl buffer (pH 7.6) containing 150 mM NaCl, 1 mM CaCl2, and 1 mM MgCl2 (TBS) for 10 min each time using an orbital shaker (KM-2; Edmund Bühler, Germany).

The locations of colonies on plate-grown bacterial cultures were documented by using a digital camera before the colonies were transferred by colony lift to circular nitrocellulose membranes (Protran BA 85; Schleicher & Schuell). The position and orientation of membranes on the agar plates was marked to allow later realignment. The membranes, after being peeled off the surface of the agar, were vigorously washed with TBS at least three times for 5 min each time to remove visible residues of transferred colonies.

Washed membranes were either left untreated or were incubated for 1 h in 15% formaldehyde to denature bacterial adhesins. Formaldehyde-treated membranes were washed three times for 5 min each time with TBS and then blocked in TBS containing 5% bovine serum albumin (fraction V; Sigma) for 2 h. The membranes were then overlaid with a suspension of fluorescein-labeled bacteria in blocking buffer at a final concentration of 2.5 × 107 organisms per ml for 45 min at 4°C in the dark in order to allow for binding to occur. After overlay, the membranes were rinsed briefly in TBS containing 0.05% Tween 20 (Bio-Rad) and then washed in the same buffer two times for 5 min to remove nonadherent bacteria. Blots were allowed to dry before the fluorescent signals of adherent bacteria were recorded by using a Typhoon 9200 imaging system (Amersham Biosciences, Freiburg, Germany).

Isolation of colonies and determination of coadhesion phenotype.

Coadhesion-positive and -negative colonies were localized by matching the position of colonies on a given plate with the image from the respective fluorescence scan. The photographs taken from the original culture plates were used as help for orientation. Colonies of interest were picked using a sterile loop and streaked out on a fresh culture plate. After 48 h of growth at 37°C and photographic documentation of the colony pattern, an additional round of colony lift and coadhesion assay was performed as described above. At this stage, the majority of colonies was expected to match the coadhesion phenotype of the originally selected colony. Singular colonies were then picked and plated out again on fresh culture plates in serial 10-fold dilutions. After another 48 h of growth, the coadhesion assay was repeated using plates on which colonies were sufficiently spaced apart to avoid cross-contamination. Normally, after two rounds of restreaking and reprobing, all of the colonies uniformly expressed the same coadhesion phenotype as the originally picked colony. At that point, the colonies were subjected to 16S rRNA gene analysis for identification and verification of purity of the respective isolates which were then stored at −80°C using the Cryobank system (see “Bacterial culture,” above).

Identification of bacterial isolates by 16S rRNA gene analysis.

Molecular species identification was carried out via sequencing of the 16S rRNA gene. DNA from bacteria was prepared by using the QIAamp DNA minikit (Qiagen, Hilden, Germany) according to the manufacturer's instructions. The almost complete 16S rRNA gene was amplified by using Roche PCR-Master premixed PCR reagent (Roche Diagnostics, Mannheim, Germany) with panbacterial primers as previously described (20). The sequences of the amplified fragments were determined on an Applied Biosystems 310 genetic analyzer using a BigDye Terminator v1.1 cycle sequencing kit (Applied Biosystems, Darmstadt, Germany) with amplification primers and additional internal sequencing primers (20) to facilitate complete double-stranded coverage of the fragments. Obtained sequence data were analyzed by using the Human Oral Microbiome Database (21) as a reference to achieve species identification.

Fluid-phase coaggregation.

Coaggregation assays were performed as previously described (22) with bacteria harvested from overnight BHI broth cultures. Different bacteria were washed, adjusted to equivalent cell densities (OD600 ~ 2.0) in coaggregation buffer, and added in equal volumes (0.5 ml) to glass tubes (10 cm by 0.75 cm). The tubes were vortex mixed for approximately 30 s, incubated briefly to allow settling of coaggregates, and scored from “+++” for maximum coaggregation to “−” for no coaggregation (6).

Nucleotide sequence accession numbers.

The sequences obtained are available under GenBank accession numbers KM225733 to KM225761 (see Tables 1 and and22 for more information).

TABLE 1
Coadhesion partners of A. naeslundii WVU45 isolated from dental plaque
TABLE 2
Coadhesion partners of S. oralis 34 isolated from dental plaque

RESULTS

Validation of the bacterial overlay technique for detecting interbacterial adhesion.

In a first set of experiments, laboratory strains of streptococci and actinomyces were used to find out whether the bacterial overlay technique can be adopted to detect interbacterial adhesive interactions. For this, A. naeslundii WVU45 and S. oralis 34, as well as S. gordonii 38, were chosen based on their well-characterized adhesive properties in the fluid-phase coaggregation assay (Fig. 1). When serial dilutions of S. oralis 34 were immobilized and overlaid with a suspension of fluorescein-labeled A. naeslundii WVU45, fluorescent signals titrated out extensively, indicating that strong interbacterial adhesion had occurred (Fig. 2A). Fewer than 5 × 105 streptococcal organisms, immobilized within the area of one well (19.6 mm2), could be detected by this technique. Based on the area theoretically occupied by an average streptococcus (ca. 0.8 μm2), we estimated that it would take roughly 107 streptococcal cells to fully cover the surface area of one dot on the nitrocellulose membrane available for the immobilization of bacteria. In other words, the detection limit of 5 × 105 cells in our assay means that bacterial coadhesion partners can still be detected, if only 2% of the available surface area is occupied by bacterial cells. Binding of the same A. naeslundii strain to immobilized RPS-deficient mutant strain S. oralis 34M was found to be weaker by a factor of ~4. A similar reduction in binding was seen when the protein adhesin on suspended strain A. naeslundii WVU45 was denatured by treatment with formaldehyde. When the putative protein adhesin on immobilized S. oralis strains was denatured by formaldehyde, binding to RPS-deficient mutant strain 34M was abolished, but binding to wild-type strain 34 persisted. Binding to the latter was only abolished when also protein adhesins on A. naeslundii WVU45 were denatured by formaldehyde.

FIG 1
Schematic diagram of cell surface proteinaceous adhesins (red) and complementary glycan receptors (black) that mediate coaggregation between the bacterial strains used in the present study. Adhesins associated with type 2 fimbriae of A. naeslundii WVU45 ...
FIG 2
Bacterial coadhesion assays performed with strains of A. naeslundii (WVU45 and WVU45M), S. oralis (34 and 34M), and S. gordonii (38 and XC3) that have known coaggregation properties (see Fig. 1). (A) Serial 2-fold dilutions of streptococcal strains 34, ...

When immobilized S. gordonii wild-type strain 38 was overlaid with a suspension of strain A. naeslundii WVU45, strong interbacterial binding was observed, paralleling the interaction seen with S. oralis 34 (Fig. 2A). No binding was observed to immobilized RPS-deficient mutant strain S. gordonii XC3. Binding to immobilized S. gordonii 38 was abolished when suspended strain A. naeslundii WVU45 was formaldehyde treated but not when immobilized strain 38 was formaldehyde treated.

Analogous interbacterial adhesion as illustrated in Fig. 1A occurred when the assay was performed in the reversed orientation, i.e., when the actinomyces strains were immobilized and overlaid by fluorescein-labeled suspensions of the streptococcal strains (Fig. 2B). Coadhesion of strains occurred with similar strengths, as in the reversed orientation (see Fig. 2A for comparison).

Overall, the observed binding phenomena replicated what had been reported earlier using the in-suspension coaggregation assay. All of the adhesive interbacterial interactions observed could be explained by the presence or absence of proteinaceous lectin-like adhesins and complementary glycan receptors on the respective strains (Fig. 1).

Detection of interbacterial binding on colony lifts from plate monocultures, using well-characterized laboratory strains.

In a follow-up set of experiments, the same laboratory strains of actinomyces and streptococci were used to test whether the overlay method can also detect interbacterial adhesion on colony lifts from plate-grown monocultures of respective partner strains (Fig. 3). Thus, when agar-grown cultures of S. oralis 34 were colony lifted and overlaid with suspensions of fluorescein-labeled A. naeslundii WVU45, strong interbacterial adhesion was observed (Fig. 3A). Interestingly, a diffuse dark halo-like effect was observed around the locations of colonies. This suggested that the agar surrounding the colonies may have been infiltrated, possibly by shedding of RPS from S. oralis 34. Weaker interbacterial binding and absence of the halo-like effect was noted when colony lifts of RPS-deficient S. oralis mutant strain 34M were overlaid with A. naeslundii WVU45. Formaldehyde treatment of the actinomyces reduced binding, and no halo effect was observed in consequence. When colony lifts of S. gordonii wild-type strain 38 were overlaid with suspensions of A. naeslundii WVU45, strong binding was observed. No binding was seen to colony lifts of RPS-deficient mutant strain XC3. Formaldehyde treatment of colony lifts of strain S. oralis 34 reduced binding of A. naeslundii WVU45 to the colonies, but halos remained. Only when colony lifts of S. oralis 34 and suspended A. naeslundii WVU45 were both formaldehyde treated was interbacterial binding fully abolished. Taken together, these observations support the concept that the halo-like effect may be caused by adhesion of the actinomyces to shed streptococcal RPS.

FIG 3
Bacterial coadhesion assays performed on colony lifts of bacteria that have known coaggregation properties (see Fig. 1). (A) Colony lifts of plate-grown streptococci (strain 34, 34M, 38, or XC3) were incubated with fluorescein-labeled A. naeslundii WVU45, ...

In all other situations tested, the outcome of interbacterial binding was as expected (see Fig. 3, using Fig. 1 for interpretation). When the coadhesion partners were exposed to each other in the reversed orientation, i.e., when colony lifts of actinomyces strains were overlaid with S. oralis 34 (Fig. 3B), analogous interbacterial interactions could be observed, with the exception that no halo-like effect could be seen. Taken together, these data demonstrate that the principal adhesive interactions shown with immobilized bacterial suspension cultures (Fig. 2) also hold true when the immobilized bacteria were derived from colony lifts from plate cultures.

Detection of interbacterial binding on colony lifts from plate cultures of mixed oral biofilm bacteria.

To take it one step further and test whether the overlay technique can be used to probe for the occurrence of unknown coadhesion partners in mixed oral biofilm cultures, samples of dental plaque were cultured on agar plates, colony lifted, and overlaid with suspensions of fluorescein-labeled A. naeslundii WVU45. As shown in Fig. 4A, a mix of coadhesion-positive and -negative colonies could be identified by this method. One positive colony (B11) and one negative colony (B18) were picked as examples, repeatedly subcultured, and overlaid until a stable homogeneous coadhesion phenotype could be confirmed. After three cycles of subculturing and reprobing, all colonies derived from originally coadhesion-positive colony B11 remained positive, and all colonies derived from coadhesion-negative colony B18 remained negative (Fig. 4A). The isolates were then analyzed by 16S ribosomal DNA sequencing and identified as Rothia dentocariosa for clone B11 and as a Rothia sp. for clone B18, respectively.

FIG 4
Identification of coadhesion partners for A. naeslundii WVU45 from early (overnight) dental plaque of individual 1. (A) Plaque biofilm was cultured on BHI agar, lifted onto a nitrocellulose membrane, and overlaid with fluorescein-labeled A. naeslundii ...

To assign expression of proteinaceous adhesins or glycan receptors to these isolated strains, the bacteria were immobilized on nitrocellulose using the dot blot apparatus. The results for a group of representative bacterial isolates with clearly distinct coadhesive properties are presented in Fig. 4B. Isolate B4, identified as S. mitis, showed interbacterial binding with A. naeslundii WVU45 that persisted when either the actinomyces or the streptococcal strain were treated with formaldehyde. Only when both strains were treated with formaldehyde was interbacterial adhesion abolished. This result indicates not only that the adhesin on actinomyces recognizes a glycan receptor on the streptococcal isolate B4 but also that the streptococcus expresses an adhesin which recognizes a glycan component on the actinomyces. A different coadhesive pattern is shown for isolate B8, which was identified as Neisseria pharyngis. This isolate showed coadhesive interactions with actinomyces only when it remained untreated, whereas formaldehyde treatment of the actinomyces did not diminish interbacterial binding. This suggests that the neisseria isolate expresses a protein adhesin which recognizes a glycan receptor on the actinomyces. Another example is represented by isolate B11 (the same as in Fig. 4A) which was identified as a Streptococcus sp. For this isolate, adhesive interactions could only be observed when the actinomyces remained untreated. Formaldehyde treatment of the immobilized isolate strain did not diminish interbacterial binding. This indicates that the streptococcal isolate B11 carries a glycan receptor recognized by an actinomyces protein adhesin. For isolate B18 (Rothia sp.), which was a coadhesion-negative colony (see Fig. 4A), no adhesive interactions could be observed under any of the conditions tested, indicating that this is a strain that apparently does not engage in coadhesion with A. naeslundii WVU45.

To explore the range of coadhesion partners in dental plaque that exhibit interbacterial binding with A. naeslundii WVU45, a larger number of coadhesion-positive and -negative colonies were isolated after overlay with this strain (Table 1). Not in all instances could the original binding phenotype of the colony be maintained through subsequent culturing (not shown in Table 1). In 4 of the 69 isolates originally tested, the phenotype could not be maintained. This is most likely due to the fact that mixed colonies had been picked, but it could also be due to spontaneous phenotype switching during subculturing. In two of these four isolates, 16S rRNA gene analysis suggested that a mixed culture had been obtained. Overall, however, all of the isolates that were determined to be negative for coadhesion proved also to be negative when tested in the coaggregation assay. An example is the isolate identified as S. mutans (Table 1, isolate A13). Conversely, all of the isolates that tested as positive in the coaggregation assay were also positive in the coadhesion assay. Interestingly, a number of isolates showed interbacterial binding in the coadhesion assay that was not detectable by the coaggregation assay (isolates A1 to A3 and A9 to A12). Also, in several instances a halo-like effect around the colony, reminiscent of the observation shown in Fig. 3A, was observed (isolates A1, A3, A9 to A12, and B8).

The expression of protein adhesins or glycan receptors was determined by subjecting the respective isolates to the overlay assay on dot blots in combination with formaldehyde treatment. Table 1 lists the coadhering partner strains that expresse a formaldehyde-sensitive proteinaceous adhesin. The situation of a protein adhesin expressed on the actinomyces interacting with a putative RPS on the immobilized coadhesion partner (resembling the interaction of A. naeslundii WVU45 with S. gordonii 38 [Fig. 1 and Fig. 2B]) was found for isolates A7, B5, B11, C1, C7, and C12. The situation of a protein adhesin expressed on the immobilized strain interacting with a putative glycan receptor on the actinomyces (resembling the interaction of A. naeslundii WVU45 with S. oralis 34M [Fig. 1 and Fig. 2A]) was found with isolates A1 to A3, A9 to A12, B8, B10, and C13. The presence of protein adhesins on both coadhesion partners interacting with complementary glycan receptors on their respective partner strains (resembling the interaction of A. naeslundii WVU45 with S. oralis 34 [Fig. 1 and Fig. 2A]) was found for colony B4.

The variety of bacterial coadhesion partners found for actinomyces clearly expands the range of previously recognized oral strains detectable by the classic coaggregation assay. Additional species were identified that have not previously been recognized to coaggregate with actinomyces, such as Rothia dentocariosa (A7, B11, and C1) and Neisseria pharyngis (B8). Coadhesion partners for the actinomyces could be found both in early (Table 1, B and C isolates) and in mature dental plaque (Table 1, A isolates).

As a further proof of principle, dental plaque cultures were also probed with S. oralis 34 (Table 2). Notably, not only were actinomyces or other species of streptococci detected as coadhesion partners for S. oralis 34 but also strains belonging to other oral bacterial species, some of which had not been recognized to coaggregate with S. oralis 34, such as Kingella oralis (isolate S6), Rothia dentocariosa (isolate S9), and Neisseria spp. (S4 and S8). The latter did not coaggregate but showed coadhesion with S. oralis 34. Notably, this type of coadhesion could not be abolished, even when both coadhesion partners were treated with formaldehyde, which indicates that other modes of coadhesion may exist that do not depend on lectin-glycan interaction.

DISCUSSION

The present study has shown that interbacterial adhesive interactions, which can be detected by the classic coaggregation assay, are equally detectable by the solid-phase bacterial overlay technique described here. However, the latter can reveal additional interactions that are not detectable by in-suspension coaggregation. The assay also indicates which of the coadhesion partners expressed the proteinaceous (formaldehyde sensitive) adhesins and which expressed the corresponding glycan receptors (formaldehyde resistant). For this, it does not matter which of the two partner strains is immobilized and which is kept in suspension. The assay further provides semiquantitative comparisons of the avidity of interbacterial binding. Most importantly, it also works on colony lifts taken from plate-grown bacterial cultures and thus can be used to efficiently screen mixed bacterial biofilm cultures for the presence of coadhering partners.

Interbacterial adhesion between two genetically distinct bacteria can occur in two major ways. (i) If both microorganisms are suspended in the planktonic phase while interacting, it is called “coaggregation” (6). (ii) If one microorganism is immobilized on a surface and the other one is suspended, it is called “coadhesion” (23). The underlying mechanisms mediating coaggregation and coadhesion were assumed to be identical (24), but it has been questioned whether an in-suspension coaggregation assay can accurately reflect what happens in vivo, e.g., in dental plaque biofilms (23). In consequence, various types of assays have been developed to better mimic coadhesion, and these approaches have added to the present understanding of biofilm formation (15, 23, 25,30). The coadhesion assay, presented here, reveals additional interbacterial adhesive interactions that are not detectable by coaggregation. Although the specificity of the coadhesion assay resembles that of the coaggregation assay (i.e., strains that are negative in coadhesion are also negative in coaggregation), its sensitivity appears to be higher in that it revealed a wider range of putative RPS-bearing strains that included nonstreptococcal species, e.g., Rothia dentocariosa. The strength of adhesive interaction with Rothia spp., together with the frequency of isolation, suggests that this genus might play an important and as-yet-underappreciated role in dental biofilm development. Notably, Rothia spp. were also frequently detected in biofilms covering oropharyngeal prostheses (31). Additional species identified as coadhesion partners in the present study that were not known to coaggregate included Neisseria spp. and Kingella oralis.

The fact that the coadhesion assay detects a wider range of interbacterial adhesive interactions may be due to two main reasons. (i) Immobilizing bacteria to a solid support results in a clustered accumulation of receptors or adhesins, which can increase the avidity of interbacterial binding. Such increased avidity in lectin-mediated binding through multivalency has been described (32, 33). (ii) Extracellular products shed by the bacteria in a colony become transferred by the colony lift to the membrane and may include RPS or adhesins. The halo-like effects seen in the coadhesion assay on colony lifts (see Fig. 2A as an example) may be one such example illustrating the importance of shed extracellular products for interbacterial adhesion and, eventually, biofilm formation.

The power of the present coadhesion assay lies in the fact that it allows researchers to use well-known reference strains as probes to efficiently screen mixed bacterial cultures for coadhering partner organisms that express complementary adhesins or glycan receptors. Using this assay, unidentified bacterial receptor-adhesin interactions, other than the ones based on recognition of RPS by type 2 actinomyces adhesins, could also be elucidated which have long been postulated to exist (34,39). In broader sense, the coadhesion assay may be used to isolate and identify coadhesion partners from any type of mixed biofilm communities occurring in the human organism or elsewhere in nature. It could equally become an efficient tool for screening genetic libraries for adhesive phenotypes of interest.

ACKNOWLEDGMENTS

We are grateful to the late Norbert Lehn, Department of Medical Microbiology and Hygiene, University of Regensburg, for helpful advice.

This study was supported by grants DFG SFB 585/B5 (S.R.) and NIDCR DE019807 (S.R.) and by the Intramural Research Program of the NIDCR, National Institutes of Health (J.O.C.).

Footnotes

Published ahead of print 8 August 2014

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