Probing of Microbial Biofilm Communities for Coadhesion Partners Stefan Ruhl,a,b Andreas Eidt,b Holger Melzl,c Udo Reischl,c John O. Cisard
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 fluorescencebased 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 fluoresceinlabeled 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.
O
ral 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 co-
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localized 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.
Received 2 June 2014 Accepted 1 August 2014 Published ahead of print 8 August 2014 Editor: A. M. Spormann Address correspondence to Stefan Ruhl, [email protected]. Copyright © 2014, American Society for Microbiology. All Rights Reserved. doi:10.1128/AEM.01826-14
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Department of Oral Biology, School of Dental Medicine, University at Buffalo, The State University of New York, Buffalo, New York, USAa; Department of Operative Dentistry and Periodontology, Dental School,b and Institute of Clinical Microbiology and Hygiene,c University Hospital Regensburg, University of Regensburg, Regensburg, Germany; Laboratory of Cell and Developmental Biology, National Institute of Dental and Craniofacial Research, National Institutes of Health, Bethesda, Maryland, USAd
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MATERIALS AND METHODS
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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 2 for more information).
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 m2), we estimated that it would take roughly 107 streptococcal cells to fully cover the surface area of one dot on the
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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 l 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 l 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 l 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. Formaldehydetreated 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
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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. 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
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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, 34M, 38, or XC3 were immobilized on nitrocellulose membranes using a dot blot apparatus, starting on the left with 107 bacteria per dot (19.6 mm2). Membranes were incubated with fluoresceinlabeled A. naeslundii WVU45, washed, and scanned for fluorescence to detect coadhesion. (B) Serial 2-fold dilutions of A. naeslundii WVU45 or WVU45M were immobilized on nitrocellulose membranes (as described above), incubated with fluorescein-labeled S. oralis 34 or S. gordonii 38, washed, and scanned for fluorescence to detect coadhesion. Suspended and/or immobilized partner strains were either left untreated or were treated with formaldehyde (indicated by gray shading) to denature cell surface adhesins prior to coadhesion assays.
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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 bind glycan receptors (i.e., RPS) on S. oralis 34 and S. gordonii 38. Putative adhesins of S. oralis 34 (and 34M) also bind glycan receptors on A. naeslundii WVU45 (and WVU45M) (40). A. naeslundii WVU45M lacks type 2 fimbriae (16), and both S. oralis 34M (17) and S. gordonii XC3 (18) lack RPS. Strain 38 does not coaggregate with strain WVU45M, and strain XC3 does not coaggregate with strain WVU45 or strain WVU45M.
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streptococci (strain 34, 34M, 38, or XC3) were incubated with fluorescein-labeled A. naeslundii WVU45, washed and scanned for fluorescence to detect coadhesion. “Halos” appeared around colonies of RPS-producing S. oralis 34. (B) Colony lifts of plate-grown A. naeslundii WVU45 or WVU45M were incubated with fluorescein-labeled S. oralis 34, washed, and scanned for fluorescence to detect coadhesion. Suspended and/or immobilized partner strains either were left untreated or were treated with formaldehyde (indicated by gray shading) to denature cell surface adhesins prior to coadhesion assays.
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
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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
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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
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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 WVU45 to reveal fluorescence-positive colony B11 and fluorescence-negative colony B18. (B) Homogeneous plate cultures expressing a stable coadhesion phenotype obtained after repeated rounds of restreaking and reprobing. Original colonies B11 and B18 were picked, repeatedly subcultured, and overlaid with fluorescein-labeled A. naeslundii WVU45 to confirm stable uniform coadhesion phenotypes. Isolates B11 and B18 were then subjected to 16S rRNA gene sequencing for identification. (C) Detection of proteinaceous cell surface adhesins (sensitive to formaldehyde) or glycan receptors (insensitive to formaldehyde) on coadhesion partners of A. naeslundii WVU45 by dot blotting. Isolates B11 and B18 along with two others (B4 and B8) from fluorescence-positive colonies were membrane immobilized, overlaid with fluorescein-labeled A. naeslundii WVU45, washed, and scanned for fluorescence to detect coadhesion. Suspended and/or immobilized partner strains either were left untreated or were treated with formaldehyde (indicated by gray shading) to denature cell surface adhesins prior to coadhesion assays. Representative examples of the four major occurring coadhesion phenotypes were chosen for this illustration.
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
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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. 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 halolike 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 fluoresceinlabeled 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. 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,
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TABLE 1 Coadhesion partners of A. naeslundii WVU45 isolated from dental plaque Closest hit/matche: Coadhesion with strain WVU45b
Adhesin-bearing partnerc
Coaggregation with strain WVU45
Taxonomic group of isolate
GenBank accession no.
Similarity (%)d
A isolates A1 A2 A3 A7 A9 A11 A12 A13
⫹* ⫹ ⫹* ⫹ ⫹* ⫹* ⫹* –
A1 A2 A3 WVU45 A9 A11 A12
– – – ⫹⫹⫹ – – – –
Actinomyces sp. Actinomyces sp. Actinomyces sp. Rothia dentocariosa Actinomyces sp. Actinomyces sp. Actinomyces sp. Streptococcus mutans
AY008314 AF287748 AY278611 M59055 AF385553 AY008315 AB545935 AJ243965
99.1 99.1 99.3 100 98.7 99.1 98.7 100
B isolates B2 B4 B5 B8 B10 B11 B18
⫹/– ⫹ ⫹ ⫹* ⫹ ⫹ –
– ⫹ ⫹⫹ ⫹⫹ ⫹⫹⫹ ⫹⫹⫹ –
Streptococcus sp. (mitis) Streptococcus mitis Streptococcus sp. (oralis) Neisseria pharyngis Streptococcus anginosus Rothia dentocariosa Rothia sp. (mucilaginosa)
AF003929 AF003929 AY278609 AJ239281 AF104678 M59055 X95483
99.0 99.8 99.9 99.7 99.6 100 98.8
C isolates C1 C2 C7 C12 C13 C18
⫹ ⫹/– ⫹ ⫹/– ⫹/– –
⫹⫹⫹ – ⫹⫹ ⫹ ⫹⫹⫹ –
Rothia dentocariosa Streptococcus sp. (mitis) Streptococcus australis Streptococcus sp. (mitis) Streptococcus mitis Staphylococcus epidermidis
M59055 AF003929 AF184974 AF003929 AF003929 D83363
100 99.5 99.5 99.5 99.9 99.9
B4 and WVU45 WVU45 B8 B10 WVU45
WVU45 WVU45 WVU45 C13
a The A isolates were obtained from interproximal plaque (late biofilm) and the B isolates were obtained from overnight plaque (early biofilm) from individual 1. The C isolates were obtained from overnight plaque from individual 2. b *, colonies surrounded by “halos” (see, for example, S. oralis 34 in Fig. 3A). c The adhesin-bearing partner was determined by the effect of formaldehyde treatment on the coadhesion of suspended or immobilized bacteria (see, e.g., Fig. 4B). d The following sequences are available under the indicated GenBank accession numbers: KM225733 to KM225740 (isolates A1 to A3), KM225741 to KM225747 (isolates B2 to B18), and KM225748 to KM225753 (isolates C1 to C18). e Closest hits, but not exact matches, are shown in parentheses. For detailed sequence information, see the deposited GenBank entries of the respective isolates.
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 form-
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aldehyde-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
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Isolatea
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TABLE 2 Coadhesion partners of S. oralis 34 isolated from dental plaque Closest hit/matche: Coadhesion with strain 34b
Adhesin-bearing partnerc
Coaggregation with strain 34
Taxonomic group of isolate
GenBank accession no.
Similarity (%)d
S2 S3 S4 S6 S7 S8 S9 S13
⫹ ⫹ ⫹* ⫹ ⫹ ⫹ ⫹ –
S2 S3 ND S6 S7 and 34 ND 34
⫹⫹ ⫹⫹⫹ – ⫹⫹ ⫹⫹⫹ ⫹/– ⫹⫹ –
Actinomyces sp. Streptococcus sanguinis Neisseria sp. (bacilliformis) Kingella oralis Actinomyces naeslundii Neisseria sp. (bacilliformis) Rothia dentocariosa Streptococcus sanguinis
AY349367 AF003928 AY005029 L06164 X81062 AY005029 M59055 AF003928
99.1 99.7 99.4 99.9 99.6 99.4 99.9 99.7
a
Isolates of interproximal plaque (late biofilm) from individual 1 are listed. *, colonies were surrounded by “halos” (see, for example, S. oralis 34 in Fig. 3A). c The adhesin-bearing partner was determined by the effect of formaldehyde treatment on the coadhesion of suspended or immobilized bacteria (see, for example, Fig. 4B). ND, coadhesion was not abolished by the formaldehyde treatment of both partners. d Sequences are available under the following GenBank accession numbers: KM225754 to KM225761 for isolates S2 to S13. e Closest hits, but not exact matches, are shown in parentheses. For detailed sequence information, see the deposited GenBank entries of the respective isolates. b
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
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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 RPSbearing 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.
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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.).
REFERENCES
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1. Pennisi E. 2005. A mouthful of microbes. Science 307:1899 –1901. http: //dx.doi.org/10.1126/science.307.5717.1899. 2. Wade WG. 2013. The oral microbiome in health and disease. Pharmacol. Res. 69:137–143. http://dx.doi.org/10.1016/j.phrs.2012.11.006. 3. Jenkinson HF. 2011. Beyond the oral microbiome. Environ. Microbiol. 13:3077–3087. http://dx.doi.org/10.1111/j.1462-2920.2011.02573.x. 4. Consortium THMP. 2012. Structure, function and diversity of the healthy human microbiome. Nature 486:207–214. http://dx.doi.org/10 .1038/nature11234. 5. Dewhirst FE, Chen T, Izard J, Paster BJ, Tanner AC, Yu WH, Lakshmanan A, Wade WG. 2010. The human oral microbiome. J. Bacteriol. 192:5002–5017. http://dx.doi.org/10.1128/JB.00542-10. 6. Cisar JO, Kolenbrander PE, McIntire FC. 1979. Specificity of coaggregation reactions between human oral streptococci and strains of Actinomyces viscosus or Actinomyces naeslundii. Infect. Immun. 24:742–752. 7. Kolenbrander PE, Palmer RJ, Jr, Rickard AH, Jakubovics NS, Chalmers NI, Diaz PI. 2006. Bacterial interactions and successions during plaque development. Periodontol. 2000 42:47–79. http://dx.doi.org/10.1111/j .1600-0757.2006.00187.x. 8. Yoshida Y, Palmer RJ, Yang J, Kolenbrander PE, Cisar JO. 2006. Streptococcal receptor polysaccharides: recognition molecules for oral biofilm formation. BMC Oral Health 6(Suppl 1):S12. http://dx.doi.org/10 .1186/1472-6831-6-S1-S12. 9. Reardon-Robinson ME, Wu C, Mishra A, Chang C, Bier N, Das A, Ton-That H. 2014. Pilus hijacking by a bacterial coaggregation factor critical for oral biofilm development. Proc. Natl. Acad. Sci. U. S. A. 111: 3835–3840. http://dx.doi.org/10.1073/pnas.1321417111. 10. Nyvad B, Kilian M. 1990. Microflora associated with experimental root surface caries in humans. Infect. Immun. 58:1628 –1633. 11. Nyvad B, Fejerskov O. 1987. Transmission electron microscopy of early microbial colonization of human enamel and root surfaces in vivo. Scand. J. Dent. Res. 95:297–307. 12. Palmer RJ, Jr, Gordon SM, Cisar JO, Kolenbrander PE. 2003. Coaggregation-mediated interactions of streptococci and actinomyces detected in initial human dental plaque. J. Bacteriol. 185:3400 –3409. http://dx.doi .org/10.1128/JB.185.11.3400-3409.2003. 13. Prakobphol A, Murray PA, Fisher SJ. 1987. Bacterial adherence on replicas of sodium dodecyl sulfate-polyacrylamide gels. Anal. Biochem. 164:5–11. http://dx.doi.org/10.1016/0003-2697(87)90359-9. 14. Walz A, Odenbreit S, Stühler K, Wattenberg A, Meyer HE, Mahdavi J, Borén T, Ruhl S. 2009. Identification of glycoprotein receptors within the human salivary proteome for the lectin-like BabA and SabA adhesins of Helicobacter pylori by fluorescence-based 2-D bacterial overlay. Proteomics 9:1582–1592. http://dx.doi.org/10.1002/pmic.200700808. 15. Lamont RJ, Rosan B. 1990. Adherence of mutans streptococci to other oral bacteria. Infect. Immun. 58:1738 –1743. 16. Cisar JO, David VA, Curl SH, Vatter AE. 1984. Exclusive presence of lactose-sensitive fimbriae on a typical strain (WVU45) of Actinomyces naeslundii. Infect. Immun. 46:453– 458. 17. Cisar JO, Brennan MJ, Sandberg AL. 1985. Lectin-specific interaction of actinomyces fimbriae with oral streptococci, p 159 –163. In Mergenhagen SE, Rosan B (ed), Molecular basis of oral microbial adhesion. American Society for Microbiology, Washington, DC. 18. Xu DQ, Thompson J, Cisar JO. 2003. Genetic loci for coaggregation receptor polysaccharide biosynthesis in Streptococcus gordonii 38. J. Bacteriol. 185: 5419 –5430. http://dx.doi.org/10.1128/JB.185.18.5419-5430.2003. 19. Takahashi Y, Ruhl S, Yoon JW, Sandberg AL, Cisar JO. 2002. Adhesion of viridans group streptococci to sialic acid-, galactose-, and Nacetylgalactosamine-containing receptors. Oral Microbiol. Immunol. 17: 257–262. http://dx.doi.org/10.1034/j.1399-302X.2002.170409.x. 20. Wilmotte A, Van der Auwera G, De Wachter R. 1993. Structure of the 16S rRNA of the thermophilic cyanobacterium Chlorogloeopsis HTF (“Mastigocladus laminosus HTF”) strain PCC7518, and phylogenetic analysis. FEBS Lett. 317:96 –100. http://dx.doi.org/10.1016/0014-5793(93)81499-P.
21. Chen T, Yu WH, Izard J, Baranova OV, Lakshmanan A, Dewhirst FE. 2010. The Human Oral Microbiome Database: a web accessible resource for investigating oral microbe taxonomic and genomic information. Database (Oxford) 2010:baq013. http://dx.doi.org/10.1093/database/baq013. 22. Egland PG, Du LD, Kolenbrander PE. 2001. Identification of independent Streptococcus gordonii SspA and SspB functions in coaggregation with Actinomyces naeslundii. Infect. Immun. 69:7512–7516. http://dx.doi.org /10.1128/IAI.69.12.7512-7516.2001. 23. Bos R, van der Mei HC, Meinders JM, Busscher HJ. 1994. A quantitative method to study co-adhesion of microorganisms in a parallel plate flow chamber: basic principles of the analysis. J. Microbiol. Methods 20:289 – 305. http://dx.doi.org/10.1016/0167-7012(94)90053-1. 24. Kolenbrander PE, Palmer RJ, Jr, Periasamy S, Jakubovics NS. 2010. Oral multispecies biofilm development and the key role of cell-cell distance. Nat. Rev. Microbiol. 8:471– 480. http://dx.doi.org/10.1038/nrmicro2381. 25. Ellen RP, Veisman H, Buivids IA, Rosenberg M. 1994. Kinetics of lactosereversible coadhesion of Actinomyces naeslundii WVU 398A and Streptococcus oralis 34 on the surface of hexadecane droplets. Oral Microbiol. Immunol. 9:364 –371. http://dx.doi.org/10.1111/j.1399-302X.1994 .tb00287.x. 26. Nagaoka S, Hojo K, Murata S, Mori T, Ohshima T, Maeda N. 2008. Interactions between salivary Bifidobacterium adolescentis and other oral bacteria: in vitro coaggregation and coadhesion assays. FEMS Microbiol. Lett. 281:183–189. http://dx.doi.org/10.1111/j.1574-6968.2008.01092.x. 27. Bos R, van der Mei HC, Busscher HJ. 1995. A quantitative method to study co-adhesion of microorganisms in a parallel plate flow chamber. II. Analysis of the kinetics of co-adhesion. J. Microbiol. Methods 23:169 –182. 28. Aspiras MB, Kazmerzak KM, Kolenbrander PE, McNab R, Hardegen N, Jenkinson HF. 2000. Expression of green fluorescent protein in Streptococcus gordonii DL1 and its use as a species-specific marker in coadhesion with Streptococcus oralis 34 in saliva-conditioned biofilms in vitro. Appl. Environ. Microbiol. 66:4074 – 4083. http://dx.doi.org/10.1128/AEM.66.9 .4074-4083.2000. 29. Millsap KW, Bos R, Van Der Mei HC, Busscher HJ. 2000. Dot assay for determining adhesive interactions between yeasts and bacteria under controlled hydrodynamic conditions. J. Microbiol. Methods 40:225–232. http://dx.doi.org/10.1016/S0167-7012(00)00131-7. 30. Park Y, Simionato MR, Sekiya K, Murakami Y, James D, Chen W, Hackett M, Yoshimura F, Demuth DR, Lamont RJ. 2005. Short fimbriae of Porphyromonas gingivalis and their role in coadhesion with Streptococcus gordonii. Infect. Immun. 73:3983–3989. http://dx.doi.org/10.1128/IAI .73.7.3983-3989.2005. 31. Elving GJ, van der Mei HC, Busscher HJ, van Weissenbruch R, Albers FW. 2002. Comparison of the microbial composition of voice prosthesis biofilms from patients requiring frequent versus infrequent replacement. Ann. Otol. Rhinol. Laryngol. 111:200 –203. 32. Collins BE, Paulson JC. 2004. Cell surface biology mediated by low affinity multivalent protein-glycan interactions. Curr. Opin. Chem. Biol. 8:617– 625. http://dx.doi.org/10.1016/j.cbpa.2004.10.004. 33. Dam TK, Brewer CF. 2008. Effects of clustered epitopes in multivalent ligand-receptor interactions. Biochemistry 47:8470 – 8476. http://dx.doi .org/10.1021/bi801208b. 34. Mizuno J, Cisar JO, Vatter AE, Fennessey PV, McIntire FC. 1983. A factor from Actinomyces viscosus T14V that specifically aggregates Streptococcus sanguis H1. Infect. Immun. 40:1204 –1213. 35. Weiss EI, Eli I, Shenitzki B, Smorodinsky N. 1990. Identification of the rhamnose-sensitive adhesin of Capnocytophaga ochracea ATCC 33596. Arch. Oral Biol. 35(Suppl):127S–130S. 36. Weiss EI, London J, Kolenbrander PE, Kagermeier AS, Andersen RN. 1987. Characterization of lectinlike surface components on Capnocytophaga ochracea ATCC 33596 that mediate coaggregation with grampositive oral bacteria. Infect. Immun. 55:1198 –1202. 37. Cassels FJ, Fales HM, London J, Carlson RW, van Halbeek H. 1990. Structure of a streptococcal adhesin carbohydrate receptor. J. Biol. Chem. 265:14127–14135. 38. Cassels FJ, London J. 1989. Isolation of a coaggregation-inhibiting cell wall polysaccharide from Streptococcus sanguis H1. J. Bacteriol. 171:4019 – 4025. 39. Kagermeier AS, London J, Kolenbrander PE. 1984. Evidence for the participation of N-acetylated amino sugars in the coaggregation between Cytophaga species strain DR2001 and Actinomyces israelii PK16. Infect. Immun. 44:299 –305. 40. Kolenbrander PE, Williams BL. 1981. Lactose-reversible coaggregation between oral actinomycetes and Streptococcus sanguis. Infect. Immun. 33:95–102.
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 fluorescencebased 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 fluoresceinlabeled 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.
O
ral 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 co-
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localized 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.
Received 2 June 2014 Accepted 1 August 2014 Published ahead of print 8 August 2014 Editor: A. M. Spormann Address correspondence to Stefan Ruhl, [email protected]. Copyright © 2014, American Society for Microbiology. All Rights Reserved. doi:10.1128/AEM.01826-14
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Department of Oral Biology, School of Dental Medicine, University at Buffalo, The State University of New York, Buffalo, New York, USAa; Department of Operative Dentistry and Periodontology, Dental School,b and Institute of Clinical Microbiology and Hygiene,c University Hospital Regensburg, University of Regensburg, Regensburg, Germany; Laboratory of Cell and Developmental Biology, National Institute of Dental and Craniofacial Research, National Institutes of Health, Bethesda, Maryland, USAd
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MATERIALS AND METHODS
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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 2 for more information).
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 m2), we estimated that it would take roughly 107 streptococcal cells to fully cover the surface area of one dot on the
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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 l 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 l 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 l 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. Formaldehydetreated 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
Detection of Interbacterial Adhesion
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. 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
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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, 34M, 38, or XC3 were immobilized on nitrocellulose membranes using a dot blot apparatus, starting on the left with 107 bacteria per dot (19.6 mm2). Membranes were incubated with fluoresceinlabeled A. naeslundii WVU45, washed, and scanned for fluorescence to detect coadhesion. (B) Serial 2-fold dilutions of A. naeslundii WVU45 or WVU45M were immobilized on nitrocellulose membranes (as described above), incubated with fluorescein-labeled S. oralis 34 or S. gordonii 38, washed, and scanned for fluorescence to detect coadhesion. Suspended and/or immobilized partner strains were either left untreated or were treated with formaldehyde (indicated by gray shading) to denature cell surface adhesins prior to coadhesion assays.
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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 bind glycan receptors (i.e., RPS) on S. oralis 34 and S. gordonii 38. Putative adhesins of S. oralis 34 (and 34M) also bind glycan receptors on A. naeslundii WVU45 (and WVU45M) (40). A. naeslundii WVU45M lacks type 2 fimbriae (16), and both S. oralis 34M (17) and S. gordonii XC3 (18) lack RPS. Strain 38 does not coaggregate with strain WVU45M, and strain XC3 does not coaggregate with strain WVU45 or strain WVU45M.
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streptococci (strain 34, 34M, 38, or XC3) were incubated with fluorescein-labeled A. naeslundii WVU45, washed and scanned for fluorescence to detect coadhesion. “Halos” appeared around colonies of RPS-producing S. oralis 34. (B) Colony lifts of plate-grown A. naeslundii WVU45 or WVU45M were incubated with fluorescein-labeled S. oralis 34, washed, and scanned for fluorescence to detect coadhesion. Suspended and/or immobilized partner strains either were left untreated or were treated with formaldehyde (indicated by gray shading) to denature cell surface adhesins prior to coadhesion assays.
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
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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
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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
Detection of Interbacterial Adhesion
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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 WVU45 to reveal fluorescence-positive colony B11 and fluorescence-negative colony B18. (B) Homogeneous plate cultures expressing a stable coadhesion phenotype obtained after repeated rounds of restreaking and reprobing. Original colonies B11 and B18 were picked, repeatedly subcultured, and overlaid with fluorescein-labeled A. naeslundii WVU45 to confirm stable uniform coadhesion phenotypes. Isolates B11 and B18 were then subjected to 16S rRNA gene sequencing for identification. (C) Detection of proteinaceous cell surface adhesins (sensitive to formaldehyde) or glycan receptors (insensitive to formaldehyde) on coadhesion partners of A. naeslundii WVU45 by dot blotting. Isolates B11 and B18 along with two others (B4 and B8) from fluorescence-positive colonies were membrane immobilized, overlaid with fluorescein-labeled A. naeslundii WVU45, washed, and scanned for fluorescence to detect coadhesion. Suspended and/or immobilized partner strains either were left untreated or were treated with formaldehyde (indicated by gray shading) to denature cell surface adhesins prior to coadhesion assays. Representative examples of the four major occurring coadhesion phenotypes were chosen for this illustration.
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
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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. 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 halolike 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 fluoresceinlabeled 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. 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,
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TABLE 1 Coadhesion partners of A. naeslundii WVU45 isolated from dental plaque Closest hit/matche: Coadhesion with strain WVU45b
Adhesin-bearing partnerc
Coaggregation with strain WVU45
Taxonomic group of isolate
GenBank accession no.
Similarity (%)d
A isolates A1 A2 A3 A7 A9 A11 A12 A13
⫹* ⫹ ⫹* ⫹ ⫹* ⫹* ⫹* –
A1 A2 A3 WVU45 A9 A11 A12
– – – ⫹⫹⫹ – – – –
Actinomyces sp. Actinomyces sp. Actinomyces sp. Rothia dentocariosa Actinomyces sp. Actinomyces sp. Actinomyces sp. Streptococcus mutans
AY008314 AF287748 AY278611 M59055 AF385553 AY008315 AB545935 AJ243965
99.1 99.1 99.3 100 98.7 99.1 98.7 100
B isolates B2 B4 B5 B8 B10 B11 B18
⫹/– ⫹ ⫹ ⫹* ⫹ ⫹ –
– ⫹ ⫹⫹ ⫹⫹ ⫹⫹⫹ ⫹⫹⫹ –
Streptococcus sp. (mitis) Streptococcus mitis Streptococcus sp. (oralis) Neisseria pharyngis Streptococcus anginosus Rothia dentocariosa Rothia sp. (mucilaginosa)
AF003929 AF003929 AY278609 AJ239281 AF104678 M59055 X95483
99.0 99.8 99.9 99.7 99.6 100 98.8
C isolates C1 C2 C7 C12 C13 C18
⫹ ⫹/– ⫹ ⫹/– ⫹/– –
⫹⫹⫹ – ⫹⫹ ⫹ ⫹⫹⫹ –
Rothia dentocariosa Streptococcus sp. (mitis) Streptococcus australis Streptococcus sp. (mitis) Streptococcus mitis Staphylococcus epidermidis
M59055 AF003929 AF184974 AF003929 AF003929 D83363
100 99.5 99.5 99.5 99.9 99.9
B4 and WVU45 WVU45 B8 B10 WVU45
WVU45 WVU45 WVU45 C13
a The A isolates were obtained from interproximal plaque (late biofilm) and the B isolates were obtained from overnight plaque (early biofilm) from individual 1. The C isolates were obtained from overnight plaque from individual 2. b *, colonies surrounded by “halos” (see, for example, S. oralis 34 in Fig. 3A). c The adhesin-bearing partner was determined by the effect of formaldehyde treatment on the coadhesion of suspended or immobilized bacteria (see, e.g., Fig. 4B). d The following sequences are available under the indicated GenBank accession numbers: KM225733 to KM225740 (isolates A1 to A3), KM225741 to KM225747 (isolates B2 to B18), and KM225748 to KM225753 (isolates C1 to C18). e Closest hits, but not exact matches, are shown in parentheses. For detailed sequence information, see the deposited GenBank entries of the respective isolates.
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 form-
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aldehyde-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
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TABLE 2 Coadhesion partners of S. oralis 34 isolated from dental plaque Closest hit/matche: Coadhesion with strain 34b
Adhesin-bearing partnerc
Coaggregation with strain 34
Taxonomic group of isolate
GenBank accession no.
Similarity (%)d
S2 S3 S4 S6 S7 S8 S9 S13
⫹ ⫹ ⫹* ⫹ ⫹ ⫹ ⫹ –
S2 S3 ND S6 S7 and 34 ND 34
⫹⫹ ⫹⫹⫹ – ⫹⫹ ⫹⫹⫹ ⫹/– ⫹⫹ –
Actinomyces sp. Streptococcus sanguinis Neisseria sp. (bacilliformis) Kingella oralis Actinomyces naeslundii Neisseria sp. (bacilliformis) Rothia dentocariosa Streptococcus sanguinis
AY349367 AF003928 AY005029 L06164 X81062 AY005029 M59055 AF003928
99.1 99.7 99.4 99.9 99.6 99.4 99.9 99.7
a
Isolates of interproximal plaque (late biofilm) from individual 1 are listed. *, colonies were surrounded by “halos” (see, for example, S. oralis 34 in Fig. 3A). c The adhesin-bearing partner was determined by the effect of formaldehyde treatment on the coadhesion of suspended or immobilized bacteria (see, for example, Fig. 4B). ND, coadhesion was not abolished by the formaldehyde treatment of both partners. d Sequences are available under the following GenBank accession numbers: KM225754 to KM225761 for isolates S2 to S13. e Closest hits, but not exact matches, are shown in parentheses. For detailed sequence information, see the deposited GenBank entries of the respective isolates. b
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
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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 RPSbearing 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.
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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.).
REFERENCES
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1. Pennisi E. 2005. A mouthful of microbes. Science 307:1899 –1901. http: //dx.doi.org/10.1126/science.307.5717.1899. 2. Wade WG. 2013. The oral microbiome in health and disease. Pharmacol. Res. 69:137–143. http://dx.doi.org/10.1016/j.phrs.2012.11.006. 3. Jenkinson HF. 2011. Beyond the oral microbiome. Environ. Microbiol. 13:3077–3087. http://dx.doi.org/10.1111/j.1462-2920.2011.02573.x. 4. Consortium THMP. 2012. Structure, function and diversity of the healthy human microbiome. Nature 486:207–214. http://dx.doi.org/10 .1038/nature11234. 5. Dewhirst FE, Chen T, Izard J, Paster BJ, Tanner AC, Yu WH, Lakshmanan A, Wade WG. 2010. The human oral microbiome. J. Bacteriol. 192:5002–5017. http://dx.doi.org/10.1128/JB.00542-10. 6. Cisar JO, Kolenbrander PE, McIntire FC. 1979. Specificity of coaggregation reactions between human oral streptococci and strains of Actinomyces viscosus or Actinomyces naeslundii. Infect. Immun. 24:742–752. 7. Kolenbrander PE, Palmer RJ, Jr, Rickard AH, Jakubovics NS, Chalmers NI, Diaz PI. 2006. Bacterial interactions and successions during plaque development. Periodontol. 2000 42:47–79. http://dx.doi.org/10.1111/j .1600-0757.2006.00187.x. 8. Yoshida Y, Palmer RJ, Yang J, Kolenbrander PE, Cisar JO. 2006. Streptococcal receptor polysaccharides: recognition molecules for oral biofilm formation. BMC Oral Health 6(Suppl 1):S12. http://dx.doi.org/10 .1186/1472-6831-6-S1-S12. 9. Reardon-Robinson ME, Wu C, Mishra A, Chang C, Bier N, Das A, Ton-That H. 2014. Pilus hijacking by a bacterial coaggregation factor critical for oral biofilm development. Proc. Natl. Acad. Sci. U. S. A. 111: 3835–3840. http://dx.doi.org/10.1073/pnas.1321417111. 10. Nyvad B, Kilian M. 1990. Microflora associated with experimental root surface caries in humans. Infect. Immun. 58:1628 –1633. 11. Nyvad B, Fejerskov O. 1987. Transmission electron microscopy of early microbial colonization of human enamel and root surfaces in vivo. Scand. J. Dent. Res. 95:297–307. 12. Palmer RJ, Jr, Gordon SM, Cisar JO, Kolenbrander PE. 2003. Coaggregation-mediated interactions of streptococci and actinomyces detected in initial human dental plaque. J. Bacteriol. 185:3400 –3409. http://dx.doi .org/10.1128/JB.185.11.3400-3409.2003. 13. Prakobphol A, Murray PA, Fisher SJ. 1987. Bacterial adherence on replicas of sodium dodecyl sulfate-polyacrylamide gels. Anal. Biochem. 164:5–11. http://dx.doi.org/10.1016/0003-2697(87)90359-9. 14. Walz A, Odenbreit S, Stühler K, Wattenberg A, Meyer HE, Mahdavi J, Borén T, Ruhl S. 2009. Identification of glycoprotein receptors within the human salivary proteome for the lectin-like BabA and SabA adhesins of Helicobacter pylori by fluorescence-based 2-D bacterial overlay. Proteomics 9:1582–1592. http://dx.doi.org/10.1002/pmic.200700808. 15. Lamont RJ, Rosan B. 1990. Adherence of mutans streptococci to other oral bacteria. Infect. Immun. 58:1738 –1743. 16. Cisar JO, David VA, Curl SH, Vatter AE. 1984. Exclusive presence of lactose-sensitive fimbriae on a typical strain (WVU45) of Actinomyces naeslundii. Infect. Immun. 46:453– 458. 17. Cisar JO, Brennan MJ, Sandberg AL. 1985. Lectin-specific interaction of actinomyces fimbriae with oral streptococci, p 159 –163. In Mergenhagen SE, Rosan B (ed), Molecular basis of oral microbial adhesion. American Society for Microbiology, Washington, DC. 18. Xu DQ, Thompson J, Cisar JO. 2003. Genetic loci for coaggregation receptor polysaccharide biosynthesis in Streptococcus gordonii 38. J. Bacteriol. 185: 5419 –5430. http://dx.doi.org/10.1128/JB.185.18.5419-5430.2003. 19. Takahashi Y, Ruhl S, Yoon JW, Sandberg AL, Cisar JO. 2002. Adhesion of viridans group streptococci to sialic acid-, galactose-, and Nacetylgalactosamine-containing receptors. Oral Microbiol. Immunol. 17: 257–262. http://dx.doi.org/10.1034/j.1399-302X.2002.170409.x. 20. Wilmotte A, Van der Auwera G, De Wachter R. 1993. Structure of the 16S rRNA of the thermophilic cyanobacterium Chlorogloeopsis HTF (“Mastigocladus laminosus HTF”) strain PCC7518, and phylogenetic analysis. FEBS Lett. 317:96 –100. http://dx.doi.org/10.1016/0014-5793(93)81499-P.
21. Chen T, Yu WH, Izard J, Baranova OV, Lakshmanan A, Dewhirst FE. 2010. The Human Oral Microbiome Database: a web accessible resource for investigating oral microbe taxonomic and genomic information. Database (Oxford) 2010:baq013. http://dx.doi.org/10.1093/database/baq013. 22. Egland PG, Du LD, Kolenbrander PE. 2001. Identification of independent Streptococcus gordonii SspA and SspB functions in coaggregation with Actinomyces naeslundii. Infect. Immun. 69:7512–7516. http://dx.doi.org /10.1128/IAI.69.12.7512-7516.2001. 23. Bos R, van der Mei HC, Meinders JM, Busscher HJ. 1994. A quantitative method to study co-adhesion of microorganisms in a parallel plate flow chamber: basic principles of the analysis. J. Microbiol. Methods 20:289 – 305. http://dx.doi.org/10.1016/0167-7012(94)90053-1. 24. Kolenbrander PE, Palmer RJ, Jr, Periasamy S, Jakubovics NS. 2010. Oral multispecies biofilm development and the key role of cell-cell distance. Nat. Rev. Microbiol. 8:471– 480. http://dx.doi.org/10.1038/nrmicro2381. 25. Ellen RP, Veisman H, Buivids IA, Rosenberg M. 1994. Kinetics of lactosereversible coadhesion of Actinomyces naeslundii WVU 398A and Streptococcus oralis 34 on the surface of hexadecane droplets. Oral Microbiol. Immunol. 9:364 –371. http://dx.doi.org/10.1111/j.1399-302X.1994 .tb00287.x. 26. Nagaoka S, Hojo K, Murata S, Mori T, Ohshima T, Maeda N. 2008. Interactions between salivary Bifidobacterium adolescentis and other oral bacteria: in vitro coaggregation and coadhesion assays. FEMS Microbiol. Lett. 281:183–189. http://dx.doi.org/10.1111/j.1574-6968.2008.01092.x. 27. Bos R, van der Mei HC, Busscher HJ. 1995. A quantitative method to study co-adhesion of microorganisms in a parallel plate flow chamber. II. Analysis of the kinetics of co-adhesion. J. Microbiol. Methods 23:169 –182. 28. Aspiras MB, Kazmerzak KM, Kolenbrander PE, McNab R, Hardegen N, Jenkinson HF. 2000. Expression of green fluorescent protein in Streptococcus gordonii DL1 and its use as a species-specific marker in coadhesion with Streptococcus oralis 34 in saliva-conditioned biofilms in vitro. Appl. Environ. Microbiol. 66:4074 – 4083. http://dx.doi.org/10.1128/AEM.66.9 .4074-4083.2000. 29. Millsap KW, Bos R, Van Der Mei HC, Busscher HJ. 2000. Dot assay for determining adhesive interactions between yeasts and bacteria under controlled hydrodynamic conditions. J. Microbiol. Methods 40:225–232. http://dx.doi.org/10.1016/S0167-7012(00)00131-7. 30. Park Y, Simionato MR, Sekiya K, Murakami Y, James D, Chen W, Hackett M, Yoshimura F, Demuth DR, Lamont RJ. 2005. Short fimbriae of Porphyromonas gingivalis and their role in coadhesion with Streptococcus gordonii. Infect. Immun. 73:3983–3989. http://dx.doi.org/10.1128/IAI .73.7.3983-3989.2005. 31. Elving GJ, van der Mei HC, Busscher HJ, van Weissenbruch R, Albers FW. 2002. Comparison of the microbial composition of voice prosthesis biofilms from patients requiring frequent versus infrequent replacement. Ann. Otol. Rhinol. Laryngol. 111:200 –203. 32. Collins BE, Paulson JC. 2004. Cell surface biology mediated by low affinity multivalent protein-glycan interactions. Curr. Opin. Chem. Biol. 8:617– 625. http://dx.doi.org/10.1016/j.cbpa.2004.10.004. 33. Dam TK, Brewer CF. 2008. Effects of clustered epitopes in multivalent ligand-receptor interactions. Biochemistry 47:8470 – 8476. http://dx.doi .org/10.1021/bi801208b. 34. Mizuno J, Cisar JO, Vatter AE, Fennessey PV, McIntire FC. 1983. A factor from Actinomyces viscosus T14V that specifically aggregates Streptococcus sanguis H1. Infect. Immun. 40:1204 –1213. 35. Weiss EI, Eli I, Shenitzki B, Smorodinsky N. 1990. Identification of the rhamnose-sensitive adhesin of Capnocytophaga ochracea ATCC 33596. Arch. Oral Biol. 35(Suppl):127S–130S. 36. Weiss EI, London J, Kolenbrander PE, Kagermeier AS, Andersen RN. 1987. Characterization of lectinlike surface components on Capnocytophaga ochracea ATCC 33596 that mediate coaggregation with grampositive oral bacteria. Infect. Immun. 55:1198 –1202. 37. Cassels FJ, Fales HM, London J, Carlson RW, van Halbeek H. 1990. Structure of a streptococcal adhesin carbohydrate receptor. J. Biol. Chem. 265:14127–14135. 38. Cassels FJ, London J. 1989. Isolation of a coaggregation-inhibiting cell wall polysaccharide from Streptococcus sanguis H1. J. Bacteriol. 171:4019 – 4025. 39. Kagermeier AS, London J, Kolenbrander PE. 1984. Evidence for the participation of N-acetylated amino sugars in the coaggregation between Cytophaga species strain DR2001 and Actinomyces israelii PK16. Infect. Immun. 44:299 –305. 40. Kolenbrander PE, Williams BL. 1981. Lactose-reversible coaggregation between oral actinomycetes and Streptococcus sanguis. Infect. Immun. 33:95–102.
