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. Author manuscript; available in PMC: 2011 Dec 10.
Published in final edited form as: Mol Microbiol. 2010 Jun 10;77(4):841–854. doi: 10.1111/j.1365-2958.2010.07252.x

The Actinomyces oris Type 2 Fimbrial Shaft FimA Mediates Coaggregation with Oral Streptococci, Adherence to RBC and Biofilm Development

Arunima Mishra 1,*, Chenggang Wu 1,*, Jinghua Yang 2, John O Cisar 2, Asis Das 3, Hung Ton-That 1,‡
PMCID: PMC2946971  NIHMSID: NIHMS213078  PMID: 20545853

Abstract

Interbacterial interactions between oral streptococci and actinomyces and their adherence to tooth surface and the associated host cells are key early events that promote development of the complex oral biofilm referred to as dental plaque. These interactions depend largely on a lectin-like activity associated with the Actinomyces oris type 2 fimbria, a surface structure assembled by sortase (SrtC2)-dependent polymerization of the shaft and tip fimbrillins, FimA and FimB, respectively. To dissect the function of specific fimbrillins in various adherence processes, we have developed a convenient new technology for generating unmarked deletion mutants of A. oris. Here, we show that the fimB mutant, which produced type 2 fimbriae composed only of FimA, like the wild type coaggregated strongly with receptor-bearing streptococci, agglutinated with sialidase-treated RBC, and formed monospecies biofilm. In contrast, the fimA and srtC2 mutants lacked type 2 fimbriae and were non-adherent in each of these assays. Plasmidbased expression of the deleted gene in respective mutants restored adherence to wild-type levels. These findings uncover the importance of the lectin-like activity of the polymeric FimA shaft rather than the tip. The multivalent adhesive function of FimA makes it an ideal molecule for exploring novel intervention strategies to control plaque biofilm formation.

Introduction

Dental plaque-related oral diseases affect a significant fraction of the human population in all age groups and continue to pose a major health problem in the modern world. Dental plaque represents one of the most complex biofilm communities known, consisting of over 700 bacterial species (Kolenbrander et al., 2006). The development of this community begins with the initial adherence and colonization of the tooth surface by the so-called pioneer bacteria, mostly oral streptococci and Actinomyces species, which form an adhesive platform to promote the colonization of bridging bacteria, which in turn attract the late colonizers of the biofilm (Rickard et al., 2003, Kolenbrander et al., 2006). Actinomyces oris, formerly Actinomyces naeslundii genospecies 2 (Do et al., 2008, Henssge et al., 2009), is the predominant organism amongst many Actinomyces spp. known to colonize the human oral cavity in all age groups (Kamma et al., 2000, Papaioannou et al., 2009, Preza et al., 2008). A striking feature of A. oris is that it interacts with other initial colonizers such as the oral streptococci as well as the bridging Gram-negative anaerobes such as Fusobacterium nucleatum (Gibbons & Nygaard, 1970, Kolenbrander et al., 2002) and thereby appears to promote biofilm development. In vitro studies, carried out in flow cells, show that A. oris is capable of enhancing fusobacterial growth in a three-species biofilm that is composed of F. nucleatum, Streptococcus oralis and A. oris, as compared to the coculture of F. nucleatum and S. oralis alone (Periasamy et al., 2009). It is likely that A. oris provides specific adhesins, which promote fusobacterial attachment and colonization. The identification of the various adhesins that mediate multibacterial adherence among the pioneers and bridging bacteria is therefore a key step toward an improved understanding of biofilm development.

The tropism of A. oris for teeth reflects the functional activities of two antigenically distinct types of cell surface fimbriae. Type 1 fimbriae mediate initial attachment of A. oris to host salivary proline-rich proteins that coat the mineral surface (Gibbons et al., 1988), whereas type 2 fimbriae are the site of a lectin-like activity that contributes both to biofilm development and to host cell activation (McIntire et al., 1978, Cisar et al., 1979, Costello et al., 1979, Ellen et al., 1980, Sandberg et al., 1986). These interactions all involve binding of specific saccharide motifs, either GalNAcβ1-3Gal or Galβ1-3GalNAc, present in the so called receptor polysaccharides (RPS) of early colonizing oral streptococci (Cisar et al., 1995, Cisar et al., 1997) or in host cells (Ruhl et al., 2000, Ruhl et al., 1996, Stromberg & Karlsson, 1990). Importantly, the receptors on host cells such as RBC and PMN are masked by terminal α2-3 linked sialic acid. Consequently, pretreatment of these cells with sialidase, an enzyme produced by A. oris, is an essential prerequisite for type 2 fimbriae-mediated attachment. Pioneering work by Yeung and colleagues originally identified FimP and FimA as the major subunits that form the structure of type 1 and 2 fimbriae, respectively (Yeung et al., 1998, Yeung & Ragsdale, 1997). Intriguingly, the primary sequences of FimP and FimA harbor the C-terminal cell wall sorting signals (CWSS), which consist of the LPXTG motif, followed by a hydrophobic domain and a positively charged tail (Ton-That et al., 2004a). This feature has led to the discovery that Gram-positive bacteria utilize a sortase-catalyzed polymerization mechanism to assemble pilus/fimbrial structures as first demonstrated in Corynebacterium diphtheriae (Ton-That & Schneewind, 2003).

The genome of A. oris MG1, a clinical isolate from a patient with gingivitis (Delisle et al., 1978), harbors two fimbrial gene clusters: the type 1 gene cluster containing fimQ-fimP-srtC1 and the type 2 cluster containing fimB-fimA-srtC2. As previously shown by immunoelectron microscopy and biochemical analysis, the type 1 fimbriae are made of FimP, which forms the fimbrial shaft and FimQ, which constitutes the fimbrial tip. Similarly, the type 2 fimbrial structure is comprised of the shaft fimbrillin FimA and the tip fimbrillin FimB (Mishra et al., 2007). We further showed that the fimbria-specific sortase SrtC2 is essential for the assembly of the type 2 fimbriae, however this sortase plays no role in the assembly of the type 1 fimbriae. It thus appears that fimbrial assembly in Actinomyces takes place by a sortase-mediated mechanism that shares many common features with that of C. diphtheriae and many other Gram-positive bacteria (Mandlik et al., 2008, Telford et al., 2006, Scott & Zahner, 2006, Marraffini et al., 2006, Proft & Baker, 2009). Consistent with this notion, when expressed in C. diphtheriae the shaft fimbrillin FimA of Actinomyces is assembled into pilus polymers by SrtD (Ton-That et al., 2004b), a corynebacterial sortase enzyme which is specific for the SpaH-type pili (Swierczynski & Ton-That, 2006). Coincidently, our phylogenetic analysis revealed that FimA is closely related to SpaH (Mishra et al., 2007), indicating their evolutionary relationship. Like SpaH, the shaft fimbrillins FimP and FimA each contain a canonical pilin motif (YPK) with a conserved lysine that likely participates in the transpeptidation reaction linking fimbrial subunits covalently via Lys-(NH-CO)-Thr isopeptide bonds. Based on these conserved features, it has been proposed that fimbria-specific sortase SrtC2 cleaves the LPXTG motif of FimA and FimB proteins between threonine and glycine and forms acyl-enzyme intermediates. Fimbrial polymerization is initiated by the formation of a heterodimer of FimB-FimA joined by the threonine residue of FimB and the lysine residue of FimA pilin motif. Continual polymerization of FimA shaft fimbrillins then ensues by continual formation of sortase-acyl-FimA intermediates and cyclic addition of FimA at the growing base of the polymer. At some point, the fimbrial polymers must be linked to the cell wall likely via the lipid II precursor (Perry et al., 2002, Ruzin et al., 2002), which remains to be demonstrated. With two types of heterodimeric fimbriae, the Actinomyces fimbrial system appears to be more closely related to that of B. cereus than C. diphtheriae (Budzik et al., 2007, Mandlik et al., 2008). However, it is unclear how the polymerization of these structures is terminated and how these structures are anchored to the cell wall. Most importantly, the precise molecular role of the shaft and tip fimbrillins in host-bacterial and inter-bacterial interactions has remained obscure.

To unravel the molecular function of specific fimbrillins, it is first necessary to engineer specific fim gene deletion mutants of Actinomyces. Here, we report the development of a new allelic-exchange technology for chromosome engineering that enabled us to generate unmarked in-frame deletion mutants of Actinomyces rapidly and efficiently. Subsequently, our studies of the deletion mutants affecting the type 2 fimbrial gene cluster have led to the surprising discovery that the shaft fimbrillin FimA is solely responsible for Actinomyces' interaction not only with S. oralis, but also its adherence to erythrocytes and even the formation of the Actinomyces biofilm. These findings open a molecular approach to understanding the role of type 2 fimbriae in biofilm formation and exploring possible novel strategies for the improved control of dental plaque formation.

Results

Development of a facile chromosome engineering system for Actinomyces oris

Although Actinomyces spp. are among the first Gram-positive bacteria in which fimbriae have been identified (Girard & Jacius, 1974), our molecular understanding of fimbrial assembly in this organism has so far been hampered due to the lack of a facile gene disruption technology. We, therefore, sought to develop an allelic exchange system in Actinomyces that can introduce a gene deletion mutation without polar effects on downstream genes. The creation of such mutants can be accomplished by a two-step process that involves integration of a plasmid at the target locus by homologous recombination, followed by plasmid excision via a second cross-over event that leaves an unmarked deletion of the target gene (Ton-That & Schneewind, 2003). Indeed in a previous study, we used the plasmid pUC19, which does not replicate in A. oris, to successfully generate such a non-polar in frame deletion of the sortase gene srtC2 (Mishra et al., 2007). However, this system proved extremely laborious not only because of low transformation efficiency and low frequency of homologous recombination in A. oris, but also because the excision event could not be selected due to the absence of a positive selection marker suitable for this organism. In some cases, screening of over 10,000 colonies did not produce a desired recombinant (Mishra et al., unpublished data). Consequently, we explored two counter-selectable markers, sacB coding for levansucrase and galK coding for galactokinase, and found galK to be suitable for Actinomyces, as described below.

Galactokinase is a key enzyme of galactose metabolism, which phosphorylates D-galactose to generate galactose 1- phosphate. The enzyme can also convert 2-deoxy-D-galactose (2-DG) into 2-deoxygalactose-1-phosphate, which, however, is not further metabolized and toxic to the cell (Reyrat et al., 1998). Thus, galK has been used as a counter-selectable marker in several bacteria, including Myxococcus xanthus (Ueki et al., 1996) and Streptococcus mutans (Merritt et al., 2007). A. oris MG1 harbors a galK homolog (gene ID ANA_0203, the A. oris MG1 genome database from www.oralgen.lanl.gov). To examine whether galK is functional in MG1, we first tested its sensitivity to 2-DG. Remarkably, when cells of MG1 (WT) were plated in the presence of 0.25% 2-DG, no colonies were observed (Fig. 1A), indicating that the Actinomyces galK gene might be suitable for counter-selection. We next went on to generate an in-frame deletion mutant of galK using a pUC19 derivative selectable with kanamycin (pCW-ΔgalK) (see Experimental Procedures). We constructed the plasmid pCW-ΔgalK that contains the 1.0 kb upstream and downstream DNA fragments flanking galK but missing the bulk of the galK cistron. This plasmid was electroporated into MG1 and its integration into the bacterial chromosome was selected with kanamycin (kan) (Fig. 1B). Like the WT strain, this kan-resistant galK-integrant (marked INT) was 2-DG sensitive, i.e. unable to grow on heart infusion agar (HIA) plates supplemented with 0.25% 2-DG (Fig 1A, compared to 1C). To obtain a galK deletion mutant, we then grew a kan-resistant colony in the absence of antibiotics and plated cells on HIA containing 0.25% 2-DG. All of the resulting DG-resistant colonies tested (12 out of 12) proved to be both kan-sensitive and galK deletion mutants (ΔgalK), as judged by colony PCR analysis. Furthermore, the introduction of a plasmid expressing galK into these ΔgalK mutants prevented the bacteria from growing in the presence of 2-DG (Fig. 1A). Thus, the ΔgalK mutant strain and the GalK-expressing plasmid together constitute a powerful system for generating gene deletion mutants of A. oris that we subsequently utilized to create specific in-frame deletion mutations in the type 2 fimbrial gene cluster containing the fimB, fimA and srtC2 genes (Fig. 1E).

Figure 1.

Figure 1

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Feasibility of GalK as a counter-selectable marker in A. oris. (A - C) Cells were streaked on heart infusion agar (HIA) plates in the absence or presence of kanamycin (kan) or 2-deoxy-D-galactose (2-DG). Shown are MG1 (WT), INT. representing integration of the suicide plasmid pCWU2 into the chromosome of MG1 strain, ΔgalK deletion mutant and its isogenic derivative expressing GalK under the control of rpsJ promoter. (D) Graphic representation of the non-replicative vector pCWU2 used to generate unmarked, in-frame gene deletions in A. oris. The kanamycin resistance cassette (KanR), GalK expression under the control of rpsJ promoter, and multiple cloning sites are indicated. (E) Graphic representation of the plasmid pΔfimB used to generate an unmarked fimB deletion mutant in the type 2 fimbrial gene cluster in the ΔgalK background.

Before proceeding to characterize the fimbrial mutants, it was important first to ascertain that the deletion of galK itself did not affect fimbrial assembly. To do so, we subjected both MG1 and its isogenic ΔgalK mutant strain to the conventional tests of fimbrial biogenesis, both microscopically and biochemically. In the first test, we treated cells with specific antibodies against individual fimbrial proteins, followed by staining with IgG-conjugated gold particles and viewed by a transmission electron microscope, as described previously (Mishra et al., 2007). Specific FimA and FimB-stained gold particles were found abundantly on the bacterial surface as well as on the fimbrial structures or the tip region (FimB), both in wild type (Fig. 2A-B) as well as in the galK deletion mutant (Fig. 2C-D). Additional electron microscopic tests showed that the deletion of galK did not affect the formation of type 1 fimbriae made of FimP and FimQ (Fig. 2; data not shown).

Figure 2.

Figure 2

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GalK is not required for the assembly of type 2 fimbriae. Wild-type A. oris and its isogenic derivative ΔgalK were immobilized on carbon grids, stained with specific antibodies against FimA (α-FimA; A and C) or FimB (α-FimB; B and D) and goat anti-rabbit IgG conjugated to 12-nm gold particles. Samples were viewed by transmission electron microscopy. Scale bars indicate the length of 0.2 μm.

We next examined fimbrial polymers in the two strains by western blotting, using normalized samples from liquid cultures grown to mid-log phase, representing the bacterial cell wall by mutanolysin treatment (Fig. 3, lanes indicated by W) and the bacteria-free culture medium (lanes marked M) (see Experimental Procedures). The culture media and the cell wall fractions were each TCA-precipitated and dissolved quantitatively in SDS-sample buffer for gel electrophoresis. Using antibodies against the major subunit FimA (α-FimA) or the minor subunit FimB (α-FimB), we observed FimA monomers (FimAM) and heterogeneous high molecular weight fimbrial polymers (HMW) in the culture medium as well as cell wall fractions obtained from the wild type strain (Fig. 3). Similar amounts of FimA/FimB polymers were also observed in the ΔgalK strain (Fig. 3; data not shown). The fact that GalK is not involved in fimbrial assembly of A. oris thus enables us to analyze next the biochemical and physiological phenotypes of specific fim gene cluster mutants, as the primary goal of this study.

Figure 3.

Figure 3

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Polymerization of the type 2 fimbriae requires fimA and is independent of galK. Culture medium (M) and cell wall (W) fractions were collected from wild type strain MG1 and its isogenic derivatives grown in mid-log phase. Equivalent protein samples were separated on 4-12% Tris-Glycine gradient gel and detected by immunoblotting with α-FimA. The positions of monomers (FimAM), high molecular-mass products (FimAHMW) of fimbrial polymerization and molecular mass markers are indicated.

FimA is necessary and sufficient for fimbrial assemly

Our previous work has shown that the assembly of type 2 fimbriae is catalyzed by sortase SrtC2 and that these fimbriae are made of FimA as the major shaft-forming subunit and FimB forming the tip region (Mishra et al., 2007). So far, it has not been possible to determine if both subunits are essential for fimbrial assembly or not. Having developed the new allele exchange technology described above, we next went on to generate both fimA and fimB deletion mutants in the ΔgalK background, using our in-frame deletion cloning vector pCWU2 that expresses galK under the control of the Actinomyces rpsJ promoter (Fig. 1D; see Experimental Procedures). We then subjected these strains to immunoblot analysis and immune-electron microscopy (IEM) described above.

The western blot in Fig. 3 shows that in both WT and ΔgalK bacteria, SrtC2 catalyzed the formation of FimA polymers (marked FimAHMW). The authenticity of these polymers is demonstrated by the fact that no α-FimA reactive polymers (or a band corresponding to the FimA monomer) were detected in the newly made ΔfimA strain (ΔfimA, lane -), and that those bands were restored upon complementation of this mutant strain with a plasmid that expresses wild type FimA. Remarkably, no apparent defect of fimbrial polymerization was observed in the fimB deletion mutant (Fig. 3). Hence, the tip protein FimB is dispensable for the formation of FimA polymers. We later showed that the assembly of the type 1 fimbriae, which are made of FimP/FimQ, is independent of type 2 fimbrial subunits (see Fig. 4).

Figure 4.

Figure 4

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Assembly of the type 2 fimbriae requires the major subunit protein FimA. Bacterial cells were immobilized on carbon grids, stained with α-FimA (A, C, E & G) or α-FimB (B, D, F & H) and goat anti-rabbit IgG conjugated to 12-nm gold particles. Samples were viewed by transmission electron microscopy. Scale bars indicate the length of 0.2 μm.

To determine whether the fimbrial polymers shown in Fig. 3 were displayed on the bacterial surface, we next analyzed intact cells microscopically. Consistent with above data, results obtained with α-FimA gold staining showed that the deletion of fimB did not affect FimA assembly or the surface display of FimA-containing fibrils (Fig. 4A-D). In control experiments, when ΔfimA cells were stained with α-FimA, no FimA-labeled gold particles were observed, while unlabeled fimbriae representing the type 1 fimbriae were clearly visible (Fig. 4E). It is important to note, however, that the ΔfimA mutant cells displayed many FimB-labeled gold particles on the bacterial surface, while none of the visible fimbrial structures (type 1 fimbriae) were labeled with α-FimB (Fig. 4F). Thus, in the absence of FimA, the tip fimbrial protein FimB itself anchored to the cell wall, a situation reminiscent of what we have observed in the various hetero-trimeric pilus systems of corynebacteria (Mandlik et al., 2007). Complementation of the ΔfimA mutant with a plasmid that expresses FimA restored the assembly of FimA fibrils and also the localization of FimB at the tip, as expected (Fig. 4G-H, compared to Fig. 2A-B).

Together, these results demonstrate that the sortase mediated assembly of the type 2 fimbrial structures requires the major subunit FimA and that FimB is dispensable for the formation of these appendages.

FimA mediates recognition of RPS-bearing oral streptococci and sialidase-treated host cells

The coaggregation of A. oris and A. naeslundii strains with receptor polysaccharides (RPS)-bearing streptococci such as S. oralis 34 and the agglutination of Actinomyces with sialidase-treated RBC both depend on lectin-like binding of Actinomyces type 2 fimbriae to Gal/GalNAc-containing receptors. To gain insight into the molecular basis of this interaction, we mixed washed cell suspensions of wild type MG1 and each mutant Actinomyces construct with equal volumes of receptor-positive or receptor-negative streptococcal or RBC suspensions in 24-well plates (see Experimental Procedures) and then examined the wells for coaggregation or hemagglutination to assess type 2 fimbriae-mediated adhesion (Fig. 5). To correlate the results from adhesion assays with cell surface antigen production, we compared wild type and mutant Actinomyces by dot immunoblotting performed with specific anti-FimA and anti-FimB antibodies as well as with an antibody against type 1 fimbriae to provide a control for equivalent spotting of nitrocellulose membranes with different Actinomyces cell suspensions.

Figure 5.

Figure 5

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FimA is required for the receptor-mediated coaggregation of Actinomyces and S. oris and for hemagglutination of red blood cells. Strains were examined for coaggregation with RPS-bearing S. oralis 34 (So 34) and RPS-negative S. oralis OC1 (So OC1), bacteria-mediated hemagglutination of untreated and sialidase-treated RBC and for reactivity with α-FimA, α-FimB and α-Type 1 pili by dot immunoblotting. The latter antibody, which was included as a spotting control, reacted similarly with all strains.

The cell surface phenotypes of wild type (WT) A. oris MG1 and the ΔgalK mutant were indistinguishable (Fig. 5). Both strains coaggregated strongly with S. oralis 34 and failed to coaggregate with S. oralis OC1, an RPS-negative isogenic mutant (Yoshida et al., 2006). Similarly, both strains agglutinated with sialidase-treated RBC but not with untreated RBC. These strains also produced comparable amounts of cell surface FimA and FimB as shown by results from dot immunoblotting (Fig. 5). Comparable coaggregation and hemagglutination assays showed no adhesion of the ΔsrtC2 mutant, which lacked high molecular weight type 2 fimbriae (Fig. 3). Dot immunoblotting (Fig. 5) detected cell surface FimA and FimB of this mutant, but at levels that were at least 10-fold lower than those of the wild type. Like the ΔsrtC2 mutant, the ΔfimA mutant lacked cell surface type 2 fimbriae (Fig. 4E) and was non adherent. Dot immunoblotting of the ΔfimA (Fig. 5) confirmed the absence of cell surface FimA as well as the presence of cell surface FimB seen by immunelectron microscopy (Fig. 4F). However, the anti-FimB immunoreactivity of this mutant (Fig. 5) was only about one third that of the wild type, which suggests that most of the FimB on wild type cells is specifically linked to FimA at the tips of type 2 fimbriae. In striking contrast with the ΔfimA mutant, the ΔfimB mutant was strongly adherent as indicated by its obvious coaggregation with S. oralis 34 and hemagglutination of sialidase-treated RBC. In accordance with results from immunoelectron microscopy of the ΔfimB mutant (Fig. 4A-D), which revealed fimbriae composed only of FimA, dot immunoblotting of this mutant (Fig. 5) detected cell surface FimA near the wild type level but no FimB. Importantly, the adhesive and antigenic phenotypes of the two deletion mutants (i.e. ΔsrtC2 and ΔfimA) were restored to those of the wild type by plasmid-based expression of each deleted gene. These results reveal clearly that the lectin-like activity that mediates both host-bacterial and interbacterial interactions is a property of the shaft fimbrillin FimA rather than the tip fimbrillin FimB.

FimA mediates the formation of Actinomyces biofilms in vitro

In addition to interactions with other plaque bacteria, the ability of A. oris to form a mono-species biofilm may also play a role dental plaque formation (Moelling et al., 2007, Periasamy et al., 2009). To assess the potential roles of FimA and FimB in mono-species biofilm formation, we grew the various Actinomyces variants in multi-well polystyrene plates containing heart infusion broth (HIB), either alone or supplemented with 1% sucrose. After 48 h, the culture medium was removed, wells were washed and dried and the adherent biofilm was stained with crystal violet. Biofilm production was measured semi-quantitatively by extracting the bound crystal violet with ethanol and measuring the A580 of the extract (Fig. 6). Biofilm formation was noted when wild type MG1 and the ΔgalK derivative were grown in the presence of sucrose (Fig. 6A) but not in HIB alone (data not shown). The ΔfimB mutant, expressing the shaft fimbrillin FimA, also formed a biofilm when grown in sucrose-containing media. In contrast, the ΔsrtC2 mutant as well as the ΔfimA mutant lacking FimA failed to form biofilm as measured by staining of wells by crystal violet (Fig. 6A & 6B). Comparable results were obtained when the different constructs were cultured with 25% human saliva (Fig. 6C). The fact that the ΔfimB mutant behaved like MG1 and that a FimA-expressing plasmid rescued biofilm development by the ΔfimA mutant lead us to conclude that FimA is essential for Actinomyces biofilm development in these assays.

Figure 6.

Figure 6

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Role of Actinomyces FimA in biofilm formation. Wild-type A. oris and its isogenic derivatives were grown in microtiter plates in HIB supplemented with 1% sucrose at 37°C with 5% CO2 for 48 h (A & B) or with 25% human saliva (C). (B & C) Bacterial biofilms were stained with crystal violet and quantification was performed by determining the absorbance at 580 nm. The values presented here are the means of at least three independent experiments and the error bar represents standard deviations.

Discussion

Formation of the mixed-species biofilm community referred to as dental plaque and the initiation of inflammation at surrounding sites depends on a wide range of bacterial-host and interbacterial interactions that mediate sequential colonization of the tooth surface by different oral species. Various oral streptococcal and actinomyces species play an initiator role in this process owing to their ability to adhere to the tooth surface (Nyvad & Kilian, 1987), coaggregate to each other and interact with surrounding host cells (Hsu et al., 1994). For A. oris, a prominent early colonizer, these interactions depend on two functionally distinct types of fimbriae or pili, of which the type 1 fimbriae mediate attachment of actinomyces to adsorbed salivary proline-rich proteins on the tooth surface and the type 2 fimbriae bind specific Gal/GalNAc-containing structures that occur in streptococcal RPS and host glycoconjugates (Cisar et al., 1997). While the distinct binding specificities of these fimbriae are well established, studies to identify the corresponding adhesins have been limited by uncertainties in subunit structures of both type 1 and type 2 fimbriae and by the lack of molecular methods for genetic manipulation of A. oris. To address the first obstacle, we recently demonstrated that A. oris MG1 fimbriae are composed of specific tip and shaft fimbrillins, designated FimQ and FimP for type 1 and FimB and FimA for type 2 fimbriae, respectively (Mishra et al., 2007). We have now overcome the second obstacle through development of a facile genetic system for the isolation of in-frame deletion mutants of A. oris, based on the use of galactokinase (galK) gene as a marker for counter-selection, and have used this system to isolate ΔfimA and ΔfimB mutants for studies of fimbriae production and adhesion. The results of these studies provide important new insights into both the assembly of type 2 fimbriae and the molecular basis of type 2 fimbriae-mediated adhesion and biofilm formation.

In agreement with prior studies by Yeung and colleagues on the role of FimA in strain T14V (Yeung et al., 1998), we demonstrated that FimA of strain MG1 is required for the formation of type 2 fimbriae (Fig. 2-4). Thus, deletion of fimA abolished type 2 fimbriae production and expression of this gene in trans restored production of these fimbriae to the wild type level (Fig. 5). Interestingly, FimB was displayed on the bacterial surface in the presence or absence of FimA (Fig. 2B and Fig. 4F), a phenomenon reminiscent of the minor pilins SpaB/SpaC of C. diphtheriae and attributed to the presence of the CWSS in the two pilins (Mandlik et al., 2007). However, in the absence of FimA, the amount of cell surface FimB was reduced at least three-fold (Fig. 5), thereby indicating that FimB on wild type A. oris resides primarily at the tips of FimA shafts. By contrast, the absence of FimB did not significantly reduce cell surface FimA (Fig. 5) or the formation of type 2 fimbriae composed solely of FimA (Fig. 4).

Type 2 fimbriae formation in the absence of the tip fimbrillin FimB was not particularly surprising in view of similar results from previous studies of pilus assembly by other Gram-positive bacteria (Ton-That & Schneewind, 2003, Ton-That & Schneewind, 2004, Dramsi et al., 2006, Rosini et al., 2006, Budzik et al., 2007). However, we were surprised to find that FimB was not required for type 2-fimbriae-mediated adhesion of A. oris to RPS-bearing S. oralis and sialidase-treated RBC (Fig. 5), in view of our earlier speculation that FimB might mediate these interactions (Mishra et al., 2007), based on the widespread role of minor pilins as adhesins (Mandlik et al., 2008). Importantly, the present findings associating the Gal/GalNAc-reactive lectin-like activity of type 2 fimbriae with FimA do not exclude the possibility that FimB is also a type 2 fimbrial adhesin. Indeed, FimB may well mediate the coaggregation of A. oris with other members of the dental plaque biofilm community (see a model in Fig. 7). Further studies are underway to identify coaggregation partners of A. oris MG1 that may also coaggregate with the ΔfimA mutant but not with the ΔfimB mutant of this strain.

Figure 7.

Figure 7

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Fimbria-mediated adherence of Actinomyces oris to oral streptococci, tooth surface and the associated host cells. Actinomyces binding to salivary proline-rich proteins that coat the tooth surface requires type 1 fimbriae, consisting of the fimbrial shaft FimP (grey) and tip fimbrillin FimQ (grey square). On the other hand, actinomyces adherence to receptor (RPS) bearing-streptococci and host cells requires the type 2 fimbrial shaft FimA, not the tip fimbrillin FimB (black circle). While the fimbrial shaft FimA also promotes cell to cell adherence via a sucrose-dependent mechanism, it is conceivable that the type 2 fimbria-dependent adherence of actinomyces to bridging colonizers requires FimB. Interbacterial interactions and adherence to host cells are indicated by two-way arrows with unknown fimbrial components shown by question marks.

While the function of FimB remains to be established, it is clear from the phenotype of the ΔfimB mutant that the SrtC2-mediated polymerization of FimA is sufficient not only for formation of type 2 fimbrial shafts (Fig. 3 and 4) but also for adhesion of A. oris to RPS-bearing S. oralis 34 and sialidase-treated RBC (Fig. 5). The fact that the ΔfimB mutant failed to coaggregate with RPS-negative S. oralis OC1 or agglutinate untreated RBC (Fig. 5) strongly implies specific binding of FimA to GalNAcβ1-3Gal motifs in S. oralis 34 RPS as well as Galβ1-3GalNAc termini displayed on sialidase-treated RBC. Although these interactions are calcium-dependent (McIntire et al., 1978, Ellen et al., 1980), sequence comparisons of FimA with C-type lectins have not revealed similarities between these proteins (unpublished data). The present findings support the proposal (Drobni et al., 2006) that differences in the structural specificity of type 2 fimbriae-mediated adhesion between strains of A. oris and A. naeslundii (Cisar et al., 1995, McIntire et al., 1983, Stromberg & Karlsson, 1990) are associated with differences in the FimA sequences of these bacteria. Importantly, such differences are not localized to specific regions of FimA but instead occur across the protein sequence, except for a few conserved proline-rich regions and certain conserved motifs, including the piliin motif, E-box and the CWSS that participate in fimbrial assembly (Mishra et al., 2007, Ton-That et al., 2004b). Homology and bioinformatics analysis of FimA failed to reveal any obvious features or domain structures that might be associated with receptor binding except for a CnaB-type domain near the C-terminus (unpublished data). It is noteworthy that the corynebacterial shaft pilin SpaA contains two CnaB-type domains, but this protein does not mediate adherence to pharyngeal cells (Kang et al., 2009, Mandlik et al., 2007). Instead, the adherence is mediated by the two minor pilins SpaB and SpaC (Mandlik et al., 2007). In addition to identifying specific regions of FimA that mediate receptor binding, it remains to be determined whether this activity is an intrinsic property of the FimA monomer or alternatively, whether it arises from SrtC2-dependent polymerization of FimA monomers and is thus, specific to the FimA shaft. Further studies to determine the crystal structure of FimA are underway to address these important questions.

Despite extensive previous functional studies of A. oris type 1 and type 2 fimbriae, our present understanding of the precise roles of these structures in initial adhesion, biofilm formation and the activation of host cells is fragmentary. This is evident not only from the possibility that FimB may be an unidentified adhesin but also from the surprising role of FimA in mono-species biofilm formation by A. oris (Fig. 6). While the underlying mechanisms involved in the formation of these biofilms remain to be explored, the FimA requirement for biofilm growth in the presence of sucrose suggests a possible link between type 2 fimbriae-mediated adhesion and cell surface levan production (Moelling et al., 2007). In addition, the effect of saliva in this process may indicate a type 2 fimbriae-mediated bridging of A. oris cells by high molecular weight salivary mucins. The present findings have opened a molecular approach for further studies of these possibilities as well as further in depth studies of type 1 and type 2 fimbriae structure, function and assembly. Insights gained at the molecular level from such studies should provide an improved basis for understanding the role of A. oris in the dental plaque biofilm development (Fig. 7) as well as possible new therapeutic strategies for controlling microbial colonization of teeth as well as other host tissue surfaces by Gram-positive bacteria.

Experimental Procedures

Bacterial strains, plasmids and media

Bacterial strains and plasmids used in this study are listed in Table 1. Actinomyces were grown in hearth infusion broth (HIB) or on HIB agar plates. Streptococci were grown in complex medium containing erythromycin (10 μg ml-1) for the maintenance of ermAM (Cisar et al., 1979). Escherichia coli strains were grown in Luria Broth. When needed kanamycin was added at a concentration of 50 μg ml-1. Rabbit-raised polyclonal antibodies against recombinant fimbrial proteins were previously obtained (Mishra et al., 2007). Reagents were purchased from Sigma unless indicated otherwise.

Table 1.

Bacterial strains and plasmids used

Strain or plasmid Genotype and description Reference
Strains
A. oris MG1 Type strain, expressing type 1 & 2 fimbriae (Mishra et al., 2007)
A. oris CW1 ΔgalK; an isogenic derivative of MG1 This study
A. oris AR1 ΔsrtC2; an isogenic derivative of MG1 (Mishra et al., 2007)
A. oris AR2 AR1 containing pSrtC2 This study
A. oris AR4 ΔfimA; an isogenic derivative of CW1 This study
A. oris AR5 AR4 containing pFimA This study
A. oris CW2 ΔfimB; an isogenic derivative of CW1 This study
A. oris AR6 CW2 containing pFimB This study
S. oralis 34 Wild-type strain (type 1 Gn RPS) (Abeygunawardana et al., 1989)
S. oralis OC1 ΔwchA; a RPS-negative isogenic mutant of 34 (Yoshida et al., 2006)
Plasmids
pHTT177 Derivative of pUC19, KanR (Mishra et al., 2007)
pSrtC2 pJRD215 expressing wild-type SrtC2 from MG1 (Mishra et al., 2007)
pCW-ΔgalK pHTT177 containing flanking fragments of galK This study
pFimA pJRD215 expressing wild-type FimA from MG1 This study
pFimB pJRD215 expressing wild-type FimB from MG1 This study
pCWU2 Derivative of pHTT177, expressing GalK under the control of the rpsJ promoter This study
pΔfimB pCWU2 containing flanking fragments of fimB This study
pJRD215 Actinomyces/E. coli shuttle vector, KanR (Mishra et al., 2007)
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Plasmid construction

pCW-ΔgalK – 1.0 kb fragments upstream and downstream of galK were amplified by PCR using Phusion DNA polymerase (New England Biolabs) with primer sets galKupF/galKupR and galKdnF/galKdnR, respectively (Table 2). Generated fragments were digested with EcoRI/XbaI and XbaI/HindIII, respectively, and ligated into the vector pHTT177 cut with EcoRI and HindIII. The generated plasmid pCW-ΔgalK was confirmed by colony PCR and restriction enzyme digestion.

Table 2.

Primers used in this study

Primer Sequence(a)
galKupF GGCGGAATTCCTCACACCGATGACGAGGAC
galKupR GGCGTCTAGACGTGGTCTCCTGTGATGGGTG
galKdnF GGCGTCTAGACTGCCCGGCGCACCGCTGAG
galKdnR GGCGAAGCTTGGGGGATGTCGGAGGGGTGGT
galK-F GGCGGGATCCCATGACCAACGATGCTCCCGTCTTC
galK-R GGCGAAGCTTGCGTCGTCGGGCTCCTGCGCAGT
PrpsJ-F GGCGCATATGCGCCCGAGCGCGGGGACCAGT
PrpsJ-R GGCGGCTAGCGGCGCCTAACCTCTCTTGTACTTG
fimB-A GGCGGAATTCAACCCCTTCTACTCGAACTACCG
fimB-B GGCGGGATCCGGTTGTGGTTGGTGTAGTTGTC
fimB-C GGCGGGATCCGCTCCCGCGGGATACCGGCTTG
fimB-D GGCGTCTAGACTGCGCCTGGATGGTCTTGTCG
SrtC2-Bam-F AAAGGATCCCAACACTGGCTGTCGGCT
SrtC2-Xba-R AAATCTAGACTAGAGCCTGTGAGTCCG
FimA-Xba-F AAATCTAGACACCGACCCACGCATGAA
FimA-Eco-R AAAGAATTCTCAGAGCGCGAGGTTCGC
FimB-Eco-F AAAAAGCTTTCAGTCGAGGTTGCAGTGAC
FimB-Hind-R AAAGAATTCGACCGTCCAGCTCATCCAG
FimA-A CGCGGATCCGACACGCGCTCCTACTCC
FimA-B CCCATCCACTAAACTTAAACACTTGTCCGACGGCGTCAC
FimA-C TGTTTAAGTTTAGTGGATGGGAAGCAGAACGCGAACCTCG
FimA-D CGCGGATCCGGTGCTTCTGGAGGCTCG
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(a)

Underlined are the restriction sites in the primers.

In-frame deletion cloning vector pCWU2 – Primer sets PrpsJ-F/PrpsJ-R and galK-F/galK-R (Table 2) were used to amplify, while appending NdeI and NheI or NheI and SspI sites to the fragments, the 5’ promoter sequence and UTR of the rpsJ (ana_0026) or the galK coding sequence, respectively, from A. oris MG1 chromosomal DNA. The PCR-amplified fragments were digested with NdeI and NheI or NheI and SspI, respectively, and ligated into the vector pHTT177 cut with NdeI and SspI. The generated plasmid pCWU2 was confirmed by restriction enzyme digestion and DNA sequencing.

pFimA – To construct pFimA, the 5’ promoter sequence and UTR of srtC2 was fused to the coding sequence of fimA. For PCR amplification, primer sets SrtC2-Bam-F/SrtC2-Xba-R and FimA-Xba-F/FimA-Eco-R (Table 2) were used to amplify, while appending BamHI and XbaI or XbaI and EcoRI sites for cloning purposes, the 5’ promoter sequence and UTR of srtC2 and the coding region including fimA from chromosomal DNA of A. oris MG1. The PCR-amplified fragments were cut by BamHI and XbaI or XbaI and EcoRI restriction enzymes and ligated into the BamHI and EcoRI sites of the vector pJRD215.

pFimB – Two primers FimB-eco-F and FimB-Hind-R (Table 2) were used to amplify the fimB promoter, the 5’ UTR region, and the fimB coding sequence, while appending EcoRI and HindIII sites for cloning purposes. The PCR-amplified DNA fragment was digested with EcoRI and HindIII and ligated into the EcoRI/HindIII-cut vector pJRD215.

Gene deletions in Actinomyces oris

Generation of the host strain A. oris ΔgalK (CW1) – To make an in-frame deletion ΔgalK mutant, the deletion vector pCW-ΔgalK was electroporated into A. oris MG1 and integration of the plasmid into the chromosome (via a Campbell-type recombination) was selected on HIB agar plates supplemented with 50 μg ml-1 kanamycin (kan). Kan-resistant colonies were grown in HIB without antibiotics overnight. The following day, 50 μl aliquots of 100-fold diluted cultures were plated on HIB agar plates containing 0.25% 2-deoxy-D-galactose (2-DG). After 3 day incubation in the presence of 5% CO2 at 37°C, 12 randomly chosen 2-DG resistant colonies were restreaked on 2-DG containing HIB agar plates. Deletion of galK was verified by colony PCR using primers galK-F and galK-R (Table 2). All 12 isolates were ΔgalK and kan-sensitive.

Generation of in-frame deletion mutants of fimA and fimB – In-frame deletion mutants were obtained by homologous recombination using GalK as counterselectable marker. A gene deletion cassette carrying the flanking regions of fimA or fimB was generated by cross-over PCR with two sets of primers A/B or C/D (Table 2), according to a previous protocol (Ton-That & Schneewind, 2003). The gene deletion cassettes were digested with appropriate restriction enzymes and cloned into corresponding sites of pCWU2. The generated plasmids were then introduced into A.oris ΔgalK strain by electroporation. The insertion obtained by homologous recombination of the plasmid into the chromosome was selected by growth at 37°C in the presence of kanamycin with an efficiency frequency in the range of 1.0×10-5 – 1.0×10-6. A kan-resistant colony was grown in HIB without antibiotics overnight. The following day, aliquots of 100-fold diluted cultures were plated on HIB agar plates containing 0.25% 2-DG. Twenty 2-DG resistant colonies were restreaked on HIA+2-DG plates and then checked for kan-sensitivity (100% efficiency). 2-DG resistant-colonies should undergo the second recombination event which could either complete the allelic exchange or reconstitute the wild-type genotype. Kan-sensitive colonies were screened for the expected deletion mutation by PCR amplification using primers A and D (~ 50% efficiency). Candidate deletion mutants were characterized by western blotting with specific antibodies as well as Southern hybridization analysis.

Immuno-electron microscopy

Actinomyces grown on HIB agar plates were suspended in 0.1 M NaCl and washed with phosphate buffered saline (PBS). For immunogold labeling, a drop of bacterial suspension in PBS was placed on nickel grids with formvar carbon support (Electron Microscopy Sciences), washed three times with PBS containing 2% bovine serum albumin (BSA) and blocked for 1 h in PBS with 0.1% gelatin. Fimbriae were stained with primary antibodies (1:100 for α-FimA and 1:50 for α-FimB ) diluted in PBS containing 2% BSA for 1 h. After washing, samples were treated with 12 nm gold-goat anti-rabbit IgG (Jackson ImmunoResearch) diluted 1:20 in PBS with 2% BSA for 1 h. The samples were finally washed with water before staining with 1% uranyl acetate and viewed in a JEOL JEM-1400 electron microscope.

Cell fractionation and western blotting

Overnight cultures of Actinomyces were used to inoculate mid-log phase cultures (1:50 dilution) at 37°C in HIB medium. Kanamycin was added to a final concentration of 50 μg ml-1 when necessary. All strains were grown till OD600 ~0.6. Normalized aliquots were fractionated into medium and cell pellets by centrifugation. The cell pellets were treated with mutanolysin (300 U ml-1) in SMM buffer (0.5 M sucrose, 10 mM MgCl2, and 10 mM maleate, pH 6.8) at 37°C overnight. After the mutanolysin treatment, soluble cell wall fractions were separated from the protoplasts by centrifugation. The culture medium and cell wall fractions were subjected to TCA precipitation and acetone wash. Fimbrial preparations were then boiled in SDS containing sample buffer, separated on 4–12% Tris-Glycine gradient gels (Invitrogen), subjected to immunoblotting with rabbit antisera (1:10,000 for α-FimA and 1:1000 for α-FimB), and detected with chemiluminescence.

Dot Immunoblotting

The reactivity of A. oris wild type or mutant strains with different specific antibodies was determined by dot immunoblotting (Yoshida et al., 2005). This involved the use of a Bio-Dot Microfiltration Apparatus (Bio-Rad) to spot nitrocellulose membranes with three-fold serial dilutions of standardized Actinomyces cell suspensions, starting with approximately 1 × 107 bacteria per spot. Membranes were blocked 1 hr in TBS containing 0.1% Tween-20 and 2% skim milk prior to incubation with either rabbit α-FimA serum (1/10,000), rabbit α-FimB serum (1/1000) of with 50 ng ml-1 immune IgG from rabbit antiserum (R56) against the type 1 fimbriae of A. oris T14V (Cisar et al., 1991). Membranes were washed to remove primary antibodies, incubated with peroxidase-conjugated goat anti-rabbit IgG (Bio-Rad) and developed using a metal-enhanced DAB substrate kit (Pierce Biotechnology).

Bacterial adhesion and cell surface antigen expression

Coaggregations assays were performed with wild type or mutant A. oris strains that were cultured in a previously described complex medium (Cisar et al., 1979) containing 0.2% glucose; kanamycin at 25 μg ml-1 was included for the growth of plasmid-bearing constructs. The streptococci used as coaggregation partners included receptor polysaccharide (RPS)-bearing S. oralis 34 and the wchA mutant of this strain, S. oralis OC1 (Yoshida et al., 2006), which is RPS-negative. Bacteria were harvested by centrifugation of stationary-phase cultures, washed in 0.02 M Tris-buffered (pH 7.5) saline containing 0.1 mM CaCl2 (TBS) and suspended to comparable cell densities (approximately 2 × 109 ml−1) based on measurements of turbidity made with a Klett-Summerson colorimeter. Coaggregations were performed in 24-well cluster plates by mixing 0.25 ml volumes of Actinomyces and streptococcal cell suspensions for a few minutes on a rotary shaker prior to photography of individual wells, which were illuminated by indirect light against a black background.

The assay for bacteria-mediated hemagglutination (Costello et al., 1979) was similar to that described for coaggregation except that red blood cells (RBC, blood group O+) were used in place of streptococci. Prior to use, RBC (4% packed cells/volume) were either untreated or treated for 1 hr at 37°C with 0.3 units sialidase ml-1 (i.e. Sigma type VIII C. perfringens N-acetyl neuraminidase) 0.02 M phosphate buffered (pH 7.4) saline containing 0.1 mM CaCl2 and 2 mg ml-1 bovine serum albumin then washed to remove added enzyme. Assays were setup in 24-well cluster plates by adding 0.25 ml of 1% RBC (packed cells per volume) to 0.25 ml of Actinomyces cell suspension (approximately 2 × 109 ml−1). Plates were incubated a few minutes on a rotary shaker at room temperature to promote hemagglutination then stored on ice prior to photography of individual wells above a white light box.

In-vitro Biofilm formation

Biofilm assays were performed according to the procedure described by Moelling et al. (Moelling et al., 2007). Briefly, Actinomyces strains were grown overnight in HIB at 37°C with shaking. Cultures were diluted 1:100 in HIB supplemented with 1% sucrose, and 3 ml of diluted culture was used to inoculate sterile, 12-well polystyrene plates (Corning, NY) at 37°C with 5% CO2. After 48 h, the wells were gently washed three times with 1.5 ml sterile PBS, dried overnight in an inverted position, and stained with 0.4 ml 1% crystal violet for 10 min. The unbound dye was completely removed by washing several times with water. The plates were dried for 3 h at 37°C. For quantitative analysis of biofilm production, biofilms in wells were destained with 3 ml of 95% ethanol for 1h without shaking. Optical density at 580 nm was measured from 150 μl aliquots of each well using a Tecan Infinite M1000 Reader. The assays were performed in triplicate. Control samples did not contain cells.

For saliva-conditioned biofilm, the assay was performed with 25% human saliva according to a published protocol (Periasamy et al., 2009). Briefly, saliva was collected from ten healthy individuals and pooled. After pretreated with 2.5 mM dithiothreitol at 4°C, the pooled saliva was centrifuged at 30,000 g for 20 minutes at 4°C. The cleared supernatant was diluted with 3 volumes of distilled water to produce 25% saliva, which was then filtered with 0.22 μm pore size filters and stored at -20°C in aliquots. Prior to each use, saliva was thawed and centrifuged to remove any precipitate resulting from freeze-thaw cycles. Microtiter plate wells were preconditioned with 200 μl of saliva for 1 h at room temperature. 20 μl of overnight cultures of Actinomyces, grown in HIB at 37°C, were added to each well at an equivalent concentration of 107 CFU. The plate was incubated at 37°C for 48 h with fresh saliva being replaced at 24 hr intervals. After incubation for 48 h, the wells were washed, stained with crystal violet and quantified as described above. The assay was performed in quadruplicates and repeated three times.

Acknowledgements

We thank I-Hsiu Huang, Elizabeth Rogers and members of our laboratory for their critical inputs. This work was supported by the National Institute of Dental and Craniofacial Research (NIDCR), NIH grant DE017382 to H. T-T. and by the Intramural Research Program of NIDCR to J.O.C.

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