Genetics and Molecular Biology

Molecular and Functional Analysis of the Type IV Pilus Gene Cluster in Streptococcus sanguinis SK36

Yi-Ywan M. Chen, Yi-Chien Chiang, Tzu-Ying Tseng, Hui-Yu Wu, Yueh-Ying Chen, Chia-Hua Wu, Cheng-Hsun Chiu
Ning-Yi Zhou, Editor
DOI: 10.1128/AEM.02788-18

ABSTRACT

Streptococcus sanguinis, dominant in the oral microbiome, is the only known streptococcal species possessing a pil gene cluster for the biosynthesis of type IV pili (Tfp). Although this cluster is commonly present in the genome of S. sanguinis, most of the strains do not express Tfp-mediated twitching motility. Thus, this study was designed to investigate the biological functions encoded by the cluster in the twitching-negative strain S. sanguinis SK36. We found that the cluster was transcribed as an operon, with three promoters located 5′ to the cluster and one in the intergenic region between SSA_2307 and SSA_2305. Studies using promoter-cat fusion strains revealed that the transcription of the cluster was mainly driven by the distal 5′ promoter, which is located more than 800 bases 5′ to the first gene of the cluster, SSA_2318. Optimal expression of the cluster occurred at the early stationary growth phase in a CcpA-dependent manner, although a CcpA-binding consensus is absent in the promoter region. Expression of the cluster resulted in a short hairlike surface structure under transmission electron microscopy. Deletion of the putative pilin genes (SSA_2313 to SSA_2315) abolished the biosynthesis of this structure and significantly reduced the adherence of SK36 to HeLa and SCC-4 cells. Mutations in the pil genes downregulated biofilm formation by S. sanguinis SK36. Taken together, the results demonstrate that Tfp of SK36 are important for host cell adherence, but not for motility, and that expression of the pil cluster is subject to complex regulation.

IMPORTANCE The proteins and assembly machinery of the type IV pili (Tfp) are conserved throughout bacteria and archaea, and yet the function of this surface structure differs from species to species and even from strain to strain. As seen in Streptococcus sanguinis SK36, the expression of the Tfp gene cluster results in a hairlike surface structure that is much shorter than the typical Tfp. This pilus is essential for the adherence of SK36 but is not involved in motility. Being a member of the highly diverse dental biofilm, perhaps S. sanguinis could more effectively utilize this structure to adhere to host cells and to interact with other microbes within the same niche.

INTRODUCTION

Streptococcus sanguinis is one of the early colonizers of the oral biofilm and a primary species of the oral microbial ecosystem. The presence of this microbe is generally associated with oral health (1–4). However, S. sanguinis along with other viridans streptococci may gain entrance to the bloodstream through injury, causing transient bacteremia and, potentially, infective endocarditis (5, 6).

The genome of S. sanguinis was first completed for strain SK36, an isolate from human dental plaque (7). One unique feature of this genome is the presence of a cluster of genes, most of which are homologs of genes participating in the biosynthesis of type IV pili (Tfp). Subsequent genome sequencing projects for several S. sanguinis strains confirmed that this pil cluster is commonly present in S. sanguinis strains (8). As the GC content of the pil cluster (43.7%) in SK36 is close to that of the genome (43.4%), it is uncertain whether S. sanguinis acquired this cluster via horizontal gene transfer (8). However, since S. sanguinis is the only known oral streptococcal species that harbors a pil cluster, the products of the cluster may provide advantages for the survival of S. sanguinis in its natural niches.

Tfp are flexible, surface-exposed, hairlike structures found in both bacteria and archaea (9). Much of the knowledge on the biology of Tfp was generated from studies on Gram-negative pathogens (10), in which Tfp are often critical for their pathogenic capacity. Recent advances in genomic sequencing have led to the discovery that the pil cluster is also present in Gram-positive bacteria (11, 12). Tfp in Gram-positive bacteria exhibit biological functions similar to those seen in Gram-negative bacteria, including adherence to both biotic and abiotic surfaces, surface-dependent twitching motility, and DNA uptake in naturally competent bacteria. For instance, the Tfp of Clostridium spp. participate in in vitro biofilm formation and surface-dependent twitching motility (13–17), Ruminococcus albus utilizes Tfp to interact with cellulose (18, 19), and the Tfp of Streptococcus pneumoniae bind to extracellular DNA to facilitate natural transformation (20). Different from the Gram-negative Tfp systems, Tfp that mediate motility and that govern genetic competence are generally two independent systems in Gram-positive bacteria, although the two systems may be indistinguishable morphologically. As seen in twitching-negative S. pneumoniae, the expression of the comG operon during the development of natural competence generates 2 or 3 flexible filaments per cell, with a diameter (5 to 6 nm) and a length (2 to 3 μm) similar to those of the twitching-active Tfp (20).

S. sanguinis harbors both com genes for genetic competence and a pil cluster for Tfp biosynthesis (7, 8). A study by Gurung et al. demonstrates that the pil cluster-encoded Tfp of S. sanguinis 2908 are able to generate force via the activity of PilT ATPase to drive twitching motility on the agar surface, confirming the biological activity of Tfp in S. sanguinis (8). Although S. sanguinis SK36 and S. sanguinis 2908 share most of the common pil genes, strain SK36 fails to produce long pili and to express twitching motility under the same growth conditions (8). On the other hand, similar to the pilin genes of S. sanguinis 2908 (pilE1 and pilE2), all three putative pilin genes of S. sanguinis SK36 (SSA_2313, SSA_2314, and SSA_2315) harbor sequences encoding the characteristic features of a type IVa pilin protein, including a class III leader peptide and a conserved N-terminal motif, which forms an α-helix (21), suggesting that strain SK36 is able to generate Tfp although perhaps with different functions than the Tfp of strain 2908. Thus, this study was designed to investigate whether the pil cluster in strain SK36 is expressed and to determine the morphology and biological functions of the protein products.

RESULTS

The pil cluster of S. sanguinis SK36 and its transcriptional organization.Based on the genome annotation of S. sanguinis SK36, a putative pil gene cluster, from SSA_2318 to SSA_2302, was identified (Fig. 1A). The basic characteristics of these open reading frames (ORFs) are summarized in Table S1 in the supplemental material. Briefly, BLASTP analysis indicated that SSA_2318 and SSA_2317 encode two motor ATPases for pilus extension and retraction, respectively. The cluster also harbors genes encoding a cytoplasmic membrane protein for Tfp biosynthesis (SSA_2316), 3 putative pilin proteins (SSA_2315, SSA_2314, and SSA_2313), 5 hypothetical proteins (SSA_2312, SSA_2310, SSA_2305, SSA_2304, and SSA_2303), a putative nitric oxide reductase (NorD), 3 putative pilus assembly proteins (SSA_2309, SSA_2308, and SSA_2307), and a pilin peptidase (SSA_2302). The proposed gene names, based on the nomenclature of the Tfp system of Pseudomonas aeruginosa (22), are listed in Fig. 1A.

FIG 1
FIG 1

(A) Schematic representation and transcriptional organization of the S. sanguinis SK36 pil cluster. The gene name based on BLASTP analysis is indicated within or above the gene. The SSA_tag number is listed below the gene. The locations of PCR products shown in panel B are indicated by horizontal lines below the gene map, above the SSA_tag number. The locations of the three transcription initiation sites determined by 5′ RACE are indicated by bent arrows on solid lines. The putative promoter demonstrated by the cat fusion study is indicated by a bent arrow on a dashed line. The putative termination sites are indicated by a lollipop structure. The region that was replaced with Ωkan in strain YC9 or erm in strain YC11 is indicated by vertical arrows. (B) RT-PCR analysis demonstrating the transcripts of the pil genes. The SSA_tag number or intergenic region corresponding to the PCR products is listed above the gel photograph. Lanes 1 through 3 are PCR products generated from cDNAs of S. sanguinis SK36, YC9, and YC11, respectively. M, 1-kb DNA ladder.

To determine whether these genes are cotranscribed as a single transcript, the presence of a contiguous transcript between neighboring genes was verified by reverse transcription (RT)-PCR (data not shown). Interestingly, a RT-PCR product was also detected between SSA_2302 and its 3′-flanking gene, lytB, encoding a peptidoglycan endo-β-N-acetylglucosaminidase, suggesting that lytB is part of the pil operon. To predict the presence of transcription terminators, intergenic regions with a length of >120 bp were subjected to analysis using ARNold. Two potential terminators were detected: one is located 174 bases 3′ to the stop codon of SSA_2307, with a free energy at −6.2 kcal mol−1, and the other is located 3 bases 3′ to the stop codon of lytB, with a free energy at −13.4 kcal mol−1. The analysis also found a terminator located 154 bases 3′ to the stop codon of SSA_2320, with a free energy at −9.8 kcal mol−1. Since the RT-PCR analysis (discussed above) suggested that the pil genes are cotranscribed as an operon, the presence of transcription terminators within the cluster indicated the possibility of differential termination.

To further examine the transcriptional organization of the pil cluster, we initially attempted to use Northern blot analysis, but this approach failed to yield a clear conclusion, presumably due to the low abundance of the transcript(s). To circumvent this difficulty, RT-PCR was performed with total cellular RNA isolated from wild-type S. sanguinis SK36 and two mutant derivatives of SK36, strains YC9 and YC11 (Fig. 1 and Table 1). Strain YC9 carries a polar mutation where the sequence encoding the first 303 amino acids of SSA_2318 was replaced with Ωkan. Strain YC11 possess a nonpolar mutation where the sequence encoding amino acids 17 through 555 of SSA_2318 was replaced with erm. The Ωkan cassette (23) possesses 3 transcription terminators at both ends; hence, transcription from the promoter(s) 5′ to SSA_2318 would be terminated at the insertion site of Ωkan, whereas the replacement with the nonpolar erm gene (24) would not affect transcription. In agreement with data from the terminator prediction analysis, specific RT-PCR products for the first 12 genes of the cluster (SSA_2318 to SSA_2307) were detected only in wild-type SK36 and YC11 but not YC9 (Fig. 1B). This result indicated that the first 12 pil genes were transcribed from the promoter(s) located 5′ to the cluster. On the other hand, as RT-PCR products specific for SSA_2305, SSA_2304, SSA_2303, SSA_2302, and lytB were found in all three strains, it is expected that an additional promoter(s) exists 5′ to SSA_2305 (see below). Interestingly, a contiguous transcript was detected between SSA_2305 and SSA_2307 in SK36 and YC11 but not YC9, indicating that the terminator located 3′ to SSA_2307 partially blocks transcription from the promoter(s) located 5′ to SSA_2318.

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TABLE 1

Bacterial strains and plasmids used in this study

Promoter mapping of the pil cluster by 5′ RACE.Three transcription initiation sites, located 847, 536, and 153 bases 5′ to the translation start site of SSA_2318, were identified using the 5′ rapid amplification of cDNA ends (RACE) system (Fig. 2A). The two distal 5′ transcription initiation sites, P3 and P2, mapped to a σ70 promoter sequence (5′-TTGACA-N17-TATACT) located at an appropriate distance from the transcription initiation sites and were designated ppilB-3 and ppilB-2, respectively. The proximal transcription initiation site, P1, failed to locate a typical σ70 promoter sequence. However, an extended −10 sequence (5′-GTTTGGTTCTAT-3′) that has been frequently found in Gram-positive bacteria (25) was observed 4 bases 5′ to the transcription initiation site. This promoter was designated ppilB-1.

FIG 2
FIG 2

Identification of the transcription initiation sites of the pil cluster by 5′ RACE. (A, left) Sequencing analysis of the final PCR products. The transcription initiation site is indicated by an inverted triangle. (Right) Sequence of the putative promoter 5′ to the transcription initiation site and distance of the listed sequence to the ATG codon of SSA_2318. The extended −10 element of ppilB-1 and the putative −35 and −10 elements of ppilB-2 and ppilB-3 are shaded. (B) The identity of the pil-specific transcript generated by ppilB-3 was verified by RT-PCR. (Left) Map displaying the relative locations of the three transcription initiation sites and primers used in PCR (indicated by inverted flags). (Right) PCR results. The primer pair used for each product is shown on the top of the gel photograph. C, PCR products generated from the S. sanguinis SK36 chromosome; +, RT included in the reaction mixtures; −, control reactions without RT; M, 1-kb DNA ladder.

It is unusual for a promoter to be so distantly located from the translation start site, such as ppilB-3; therefore, RT-PCR analysis was performed to verify this result. Specifically, an antisense primer of SSA_2318 (primer AS), located 26 bases 3′ to the translational start codon, was paired with two sense primers, located 9 bases 3′ (primer 3S) and 25 bases 5′ (primer 5S) to P3, respectively (Fig. 2B). Only the primer pair AS/3S was able to generate a PCR product, confirming the mapping result. This finding confirms that the expression of the pil operon includes an untranslated region of >800 bases.

Functional analysis of the putative promoters.To analyze the activities of the three promoters, the putative −10 element in one or two promoters was mutated by site-directed mutagenesis and used in the construction of promoter-chloramphenicol acetyltransferase (CAT) gene (cat) translational fusions (Fig. 3). All fusions were integrated into the chromosome of SK36 at SSA_1656, encoding a putative nisin resistance protein (NSR). SSA_1656 was selected for integration as NSR has been shown to catalyze the degradation of nisin in Lactococcus lactis (26), and thus, the loss of SSA_1656 should have no impact on the expression of the pilB promoters. Furthermore, inactivation of this locus does not affect the growth of S. sanguinis SK36 (data not shown). When comparing the CAT-specific activities in strains harboring a single pilB promoter-cat fusion (ppilB_P1, ppilB_P2, and ppilB_P3 strains), the ppilB_P3 strain exhibited the highest CAT-specific activity, compared to the ppilB_P1 and ppilB_P2 strains (Fig. 3), and the activity was close to that detected in the wild-type pilB promoter-cat fusion strain (ppilB_WT strain), indicating that ppilB-3 was the dominant promoter driving the expression of the pil cluster. In the absence of functional ppilB-3, i.e., fusions containing native ppilB-2 (ppilB_P2 strain), ppilB-1 (ppilB_P1 strain), or both (ppilB_P1+P2 strain), the CAT-specific activity was approximately 30% of that measured in the ppilB_WT strain. Moreover, no additive effects on CAT-specific activity were observed in the ppilB_P1+P3 or ppilB_P2+P3 strain, compared to the activity measured from the ppilB_P3 strain, suggesting that the three promoters do not function cooperatively.

FIG 3
FIG 3

CAT-specific activities of the recombinant S. sanguinis strains harboring a wild-type pilB promoter-cat translational fusion or fusions driven by the derivatives of the promoter region. The relative locations of the three promoters are shown at the top. The name of the fusion strain and its genotype are listed to the left of the graph describing CAT-specific activity. The location of spe in the fusion is indicated by an inverted triangle. The putative −10 element was replaced with 5′-CTGCAG (open boxes) and 5′-CTCGAC (filled boxes) by site-directed mutagenesis. All strains were harvested at an OD600 of 0.8. Values are the means and standard deviations from three independent experiments. Significant differences between the fusion of the wild-type promoter and fusions of the mutated promoters were analyzed using one-way ANOVA followed by a Dunnett test. *, P < 0.01.

As the insertion of Ωkan in SSA_2318 (strain YC9) did not block the transcription of SSA_2305 or that of further downstream ORFs (Fig. 1B), it was suggested that transcription could be initiated in the intergenic region between SSA_2307 and SSA_2305. Since a conserved ribosomal binding site (RBS) was not observed 5′ to SSA_2305, both a transcriptional and a translational cat fusion were established in S. sanguinis SK36 at SSA_1656 to examine the function of this region. In the translational fusion strain TY26, the 490-bp region 5′ to the start codon of SSA_2305 was directly fused to the cat gene, whereas the 7 bases 5′ to the BamHI fusion site were replaced with the native RBS (5′-GGAGGCA) of cat in the transcriptional fusion strain (TY25) (Fig. 4). Approximately 30 nmol mg−1 min−1 CAT-specific activity was observed in strain TY25, whereas no activity was detected in strain TY26. This result suggests that a functional promoter, designated p2305, is present 5′ to SSA_2305 but that the translation efficiency of SSA_2305 in S. sanguinis SK36 was relatively weak.

FIG 4
FIG 4

CAT-specific activities of the recombinant S. sanguinis strains harboring either a transcriptional cat (TY25) or a translational cat (TY26) fusion of p2305. The sequence 5′ to SSA_2305 in wild-type SK36 or cat in strains TY25 and TY26 is listed below the graph. Bases identical to SK36 are in gray. The RBS of cat is in italic type. The BamHI site was engineered for cloning purposes. Both strains were harvested at an OD600 of 0.8. Values are the means and standard deviations from three independent experiments. N.D., not detectable.

CcpA is required for optimal expression of the pilB promoters.While examining the protein encoded by SSA_2315 in the total lysates, including the cell wall and cytoplasmic fractions, of S. sanguinis SK36 by immunoblotting, we noticed that a larger amount of SSA_2315 protein was detected from cultures at late log phase than from cultures at early log phase (see Fig. S1 in the supplemental material), suggesting that the expression of the pil proteins was influenced by growth phase. However, extensive sequence analysis failed to identify any known consensus sequences for regulator binding in the 5′-flanking region of SSA_2318. Nevertheless, we were encouraged by a study of Clostridium perfringens (27) in which the Tfp-mediated twitching activity was shown to be regulated by catabolite control protein A (CcpA), even though the consensus binding motif for CcpA, cre, is not found in any of the Tfp biosynthesis genes. Thus, we investigated a possible role for CcpA in pil expression by examining the expression of the ppilB_WT-cat fusion in wild-type S. sanguinis SK36, its CcpA-null derivative (ΔccpA), and the ccpA-complemented strain (CΔccpA). In agreement with our observations regarding the production of SSA_2315 protein, the CAT-specific activity expressed by ppilB_WT-cat in wild-type SK36 was upregulated at early stationary phase (optical density at 600 nm [OD600] of 0.8), compared to early log phase (OD600 of 0.3). Deletion of ccpA reduced the CAT-specific activity at both growth phases; however, expression at an OD600 of 0.8 was still elevated, compared to that from cultures grown to an OD600 of 0.3 (Fig. 5). These results indicate that CcpA positively regulates the expression of the pilB promoters. To further analyze regulation by CcpA at different growth phases, the expression of ccpA in wild-type SK36 at an OD600 of 0.3 and an OD600 of 0.8 was measured by RT-PCR. The result showed that ccpA was expressed at both growth stages at a level similar to that of dnaA, suggesting that the activation of CcpA, rather than the expression of ccpA, is involved in the regulation of pilB promoters.

FIG 5
FIG 5

Impact of CcpA on expression of the wild-type pilB promoters in response to growth phase. The CAT-specific activities of wild-type S. sanguinis SK36 (WT), its ccpA deletion derivative (ΔccpA), and the complementation strain (CΔccpA) at OD600s of 0.3 (I) and 0.8 (II) are shown. Values are the means and standard deviations from three independent experiments. Significant differences between strains at each OD600 value were analyzed using one-way ANOVA followed by a Tukey test. ***, P < 0.0001; **, P < 0.001; *, P < 0.01.

The surface structure encoded by the pil cluster.To determine whether the products of the pil gene cluster are essential for the assembly of a surface structure, wild-type SK36 and its isogenic mutant strain YC6, in which the region containing genes encoding the putative pilin proteins (SSA_2313, SSA_2314, and SSA_2315) was replaced by a nonpolar erm gene, were labeled with anti-SSA_2315 antiserum and examined by transmission electron microscopy (TEM). Gold particles were observed on a short hairlike structure of approximately 150 nm on the surface of cells from cultures of strain SK36, whereas gold particles were absent in cells from cultures of strain YC6 (Fig. 6), indicating that the pil gene cluster is required for the biosynthesis of the Tfp of S. sanguinis SK36 and that the product of SSA_2315 is one of the subunits of this structure.

FIG 6
FIG 6

Piliation of S. sanguinis examined by TEM. Cells were immunogold labeled with anti-SSA_2315 antiserum and examined by TEM at a ×200,000 magnification, as detailed in Materials and Methods.

Tfp participate in adherence of S. sanguinis SK36 to mammalian cells and affect in vitro biofilm formation.To investigate whether the Tfp of S. sanguinis SK36 participate in binding to human epithelial cells, the efficiencies of adherence of S. sanguinis SK36 and its derivatives to HeLa and oral squamous cell carcinoma (SCC-4) cells were examined. Since SSA_1632, SSA_1633, and SSA_1634 are reported to participate in the binding of SK36 to HeLa cells (28), we also constructed a mutant derivative of SK36 in which the region containing these three ORFs was replaced with erm. The resulting strain, TY10, was used as a control strain. In agreement with data from the previous report, the efficiency of adherence of strain TY10 to HeLa cells was approximately 20% lower than that of wild-type SK36 (Fig. 7A), but the difference was not statistically significant in our hands (P = 0.0214). Conversely, deletion of SSA_2313 to SSA_2315 in strain YC6 resulted in an approximately 40% reduction in adherence to both HeLa and SCC-4 cells compared to SK36. A wild-type level of adherence was observed in the complementation strain TY11, in which the erm gene in YC6 was replaced with a fragment containing SSA_2313 to SSA_2315 fused to a nonpolar kan gene at the 5′ end. Additionally, the efficiency of adherence of SK36 to SCC-4 cells was approximately 1.3-fold higher than that to HeLa cells, suggesting that S. sanguinis SK36 interacts more efficiently with oral epithelial cells than with HeLa cells in a Tfp-independent manner (Fig. 7B). Nevertheless, the results demonstrate that Tfp play a major role in the adherence of S. sanguinis SK36 to host cells.

FIG 7
FIG 7

Efficiencies of adherence of S. sanguinis SK36 and its derivatives to HeLa (A) and SCC-4 (B) cells. The percent adherence was calculated as (CFU adhered/CFU of the inoculum) × 100%. The numbers are the means and standard deviations from three independent experiments. All reactions were done in triplicate. Significant differences between strains were analyzed using one-way ANOVA followed by a Tukey test. ***, P < 0.0001; **, P < 0.001; *, P < 0.01.

Deletion of the putative pilin genes (strain YC6) did not reduce biofilm formation significantly (Fig. 8). Blocking transcription from SSA_2318 to SSA_2307 (strain YC9) or from SSA_2305 to lytB (strain YC10) by a polar mutation reduced approximately 25 to 30% of the biofilm mass compared to wild-type SK36. Interestingly, deletion of the putative pilin peptidase SSA_2302 (strain YC5) led to the least biofilm formation (Fig. 8). Taken together, the data show that expression of the pil cluster was associated with optimal biofilm formation.

FIG 8
FIG 8

Biofilm formation of S. sanguinis SK36 and its derivatives. The final growth yield was measured at 490 nm (open bars), and the biofilm mass was quantified at 562 nm (filled bars). The values are the means and standard deviations from three independent experiments. Significant differences between wild-type SK36 and its derivatives were analyzed using one-way ANOVA followed by a Dunnett test. ***, P < 0.0001.

DISCUSSION

Although less is known about the functions of Tfp of Gram-positive origin than those of Tfp of Gram-negative origin, recent genomic and functional analyses have confirmed that conservation of Tfp extends to Gram-positive Firmicutes, specifically Clostridium spp. and S. sanguinis (12). Traditionally, Tfp are divided into type IVa and type IVb based on the length of the leader peptide of the pilin protein and the size of the mature pilin protein (29). Furthermore, genes involved in the biogenesis of type IVa pili are generally located in several clusters on the chromosome, whereas genes for type IVb pilus biogenesis are located in a single cluster (10). Interestingly, the pil genes of S. sanguinis, which produce type IVa pili, are clustered. In addition, Tfp-containing Gram-positive bacteria seem to use different Tfp systems for motility and for DNA uptake during natural transformation. Similarly, inactivation of the pil genes did not affect the competence of S. sanguinis SK36, suggesting that S. sanguinis SK36 possesses at least two independent Tfp systems. Thus, SSA_0642, which is annotated as PilD, is likely to participate in the processing of the pilin proteins for competence.

The presence of multiple pilin proteins in a Tfp system is not unique to S. sanguinis SK36. For instance, R. albus harbors two tandemly located pilA genes, pilA1 and pilA2. PilA1 is part of the Tfp structure and is responsible for cellulose binding, whereas PilA2 is located in the membrane fraction and is essential for the assembly of Tfp (18). The location of the three putative pilin proteins on the Tfp structure and whether all three proteins participate in the binding of S. sanguinis SK36 to host cells are currently unknown. Upon examination of the pil clusters from all sequenced S. sanguinis strains, we found that the most significant difference between clusters is the number of putative pilin genes, ranging from a single pilin gene to 3 pilin genes. Sequence analysis of these pilin genes also revealed that, with the exception of clusters containing a single pilin gene, pilin genes within a cluster generally are more homologous to each other than to pilin genes from a different cluster. A preliminary phylogenetic analysis of the pilin proteins from the twitching-positive strain 2908 and the nontwitching strains SK1 and SK36 found that the pilin protein of strain SK1 is more closely related to that of strain 2908 than to that of strain SK36. Thus, it is suggested that the twitching phenotype is not directly related to a specific gene arrangement or a specific sequence of the pilin gene(s) in S. sanguinis.

The rationale for possessing three functional promoters to drive the expression of the pil cluster is unclear. The long untranslated message resulting from the activity of ppilB-3 would suggest that the system is regulated transcriptionally or posttranscriptionally. For instance, pilA1 of Clostridium difficile is preceded by a c-di-GMP transcriptional riboswitch, and the transcription of pilA1 is upregulated by c-di-GMP (15). To test whether pil expression in S. sanguinis SK36 was regulated by a similar mechanism, attempts were made to identify potential riboswitch structures in the 5′-flanking region of SSA_2318 using various prediction tools. However, no riboswitch structure was identified. Thus, the impact of the untranslated region on pil expression remains to be determined. It is also of note that ppilB-3 and ppilB-2 possess identical −10 and −35 elements, and in both cases, the two elements are separated by 17 bases. Yet the CAT-specific activity in the strain carrying the ppilB_P3-cat fusion is 3-fold higher than that in the strain carrying the ppilB_P2-cat fusion, suggesting that the activity of these two promoters is modulated differently by the flanking sequences. Sequence analysis revealed that the 20 bases 5′ to the −35 element of ppil-3 are AT rich and contain two AAAT repeats, which may serve as an UP element to enhance transcription (30). We also cannot rule out the possibility that the activation of ppilB-2 and ppilB-1 requires specific growth conditions, as seen in Actinobacillus pleuropneumoniae, in which the pil operon is expressed only when the bacteria adhere to lung epithelial cells (31).

How CcpA modulates pil gene expression in response to growth phase in S. sanguinis SK36 is currently unclear. It is likely that CcpA regulates pil expression indirectly, similar to the regulation reported for C. difficile (32) and others (33). We also cannot rule out the possibility that an atypical cre is recognized by CcpA in S. sanguinis SK36, as seen in Streptococcus suis (34) and Clostridium acetobutylicum (35). Comparable levels of ppilB_WT-derived CAT-specific activity were observed between glucose-grown and galactose-grown cultures (data not shown), suggesting that CcpA exerts its regulation in a glucose-independent manner. On the other hand, a recent study by Ota et al. (36) also observed growth phase-dependent expression of pilT, a homolog of SSA_2317 in strain SK36, in S. sanguinis ATCC 10556. This study found that the expression of pilT in S. sanguinis ATCC 10556 is repressed by a small RNA, csRNA1, although growth-phase-dependent expression is independent of csRNA1 regulation. Thus, expression of the pil genes is likely to be regulated by additional regulators.

A study by Okahashi et al. (28) indicated that SSA_1632 (also named PilA), SSA_1633 (PilB), and SSA_1634 (PilC) together form a surface structure. This structure participates in the binding and invasion of S. sanguinis SK36 to HeLa cells and human oral epithelial cells (HSC-2). Although both Tfp and this surface structure participate in the adherence of SK36 to host cells, they are distinct structures. Specifically, downstream of SSA_1632 is a putative sortase, SrtC (SSA_1631), and all three Pil proteins contain a sortase-dependent LPXTG cell wall-anchoring motif, although neither characteristic is associated with the Tfp system. Nevertheless, we could not confidently evaluate the impact of Tfp on invasion due to the difference in the efficiencies of adherence to host cells between wild-type SK36 and strain YC6.

The activity and morphology of Tfp of S. sanguinis SK36 are quite different from those of strain 2908 (8). The nature of the difference is unknown. The disparity in the pil gene cluster between these two strains is limited to the central region of the cluster, which contains 6 genes in SK36 (SSA_2315, SSA_2314, SSA_2313, SSA_2312, norD, and SSA_2310) and 5 genes in 2908 (pilE1, pilE2, pilA, pilB, and pilC), suggesting that this region may play a major role in the morphology and motility of the Tfp. Furthermore, S. sanguinis 2908 could produce pili with only one of the PilE proteins (8), although it is unknown whether the biological functions of these two types of Tfp are the same. Whether SSA_2315, SSA_2314, and SSA_2313 are all required for the production of the hairlike structure in S. sanguinis SK36 is unknown, but the results from our biofilm study suggest that additional proteins participating in biofilm formation are processed by SSA_2302, a putative pilin peptidase (YC5 in Fig. 8).

In conclusion, this study demonstrates that the expression of the pil cluster in S. sanguinis SK36 is regulated by growth phase. The protein products of the cluster form a surface short hairlike structure that binds to host epithelial cells and modulates biofilm formation. The hairlike structure is much shorter than the known Tfp from both Gram-positive and -negative systems, although genes within this cluster share significant homology with the known pil genes. The biological function of Tfp likely provides an advantage for S. sanguinis SK36 to establish infection in the oral cavity.

MATERIALS AND METHODS

Bacterial strains and culture conditions.All strains and primers used in this study are listed in Tables 1 and 2, respectively. S. sanguinis strains were cultivated in Todd-Hewitt (TH) broth at 37°C in 10% CO2. Where indicated, erythromycin (Em) at 10 μg ml−1, kanamycin (Km) at 250 μg ml−1, or spectinomycin (Sp) at 200 μg ml−1 was included for selecting recombinant S. sanguinis strains. For maintaining recombinant Escherichia coli strains, ampicillin (Ap) at 100 μg ml−1 or Sp at 200 μg ml−1 was used.

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TABLE 2

Primers used in this study

In silico analysis of the pil cluster.The sequence of the pil cluster was extracted from the genome of S. sanguinis SK36 (GenBank accession number NC_009009.1). The basic characteristics of all ORFs were analyzed using Vector NTI advance 11 (Invitrogen). The presence of potential intrinsic transcription terminators was analyzed by using ARNold (http://rna.igmors.u-psud.fr/toolbox/arnold/index.php).

RNA isolation, RT-PCR, and 5′ RACE.Total cellular RNA was isolated from S. sanguinis according to a method described previously by Chen et al. (37) and further purified using an RNeasy purification kit (Qiagen). To determine whether a contiguous transcript exists between the neighboring genes by RT-PCR, cDNA was generated from 2 µg of purified RNA using avian myeloblastosis virus (AMV) reverse transcriptase and random primers (Promega). Five percent of the generated cDNA was then used in a PCR. The transcriptional start site(s) of the pil cluster was determined using the 5′ RACE system (Invitrogen). Five micrograms of total cellular RNA from S. sanguinis SK36 was used in a RACE reaction. cDNA was generated from total RNA using three SSA_2318-specific primers, pil19960S, pil19730S, and pil19810S, which contain the antisense sequence of SSA_2318 and are located 226, 456, and 376 bases 3′ to the translation start site of SSA_2318, respectively. RNA was isolated from three independent cultures, and samples from each culture were used in the RACE analysis. After the addition of the poly(C) tail to the cDNA pieces, the abridged anchor primer (AAP) was paired with primers pil20050S, pil19845S, and pil19890S to amplify the pil-specific cDNA. A total of 0.1% of the resulting PCR product was reamplified by nested PCR using primer pil20180S and the abridged universal amplification primer (AUAP). The final PCR products were separated by gel electrophoresis and purified individually for sequencing analysis.

Construction of recombinant S. sanguinis strains.All recombinant strains of S. sanguinis SK36 were generated by ligation mutagenesis (38). Briefly, two DNA fragments, 5′ and 3′ to the target site, respectively, were amplified from S. sanguinis SK36 by specific primers. The PCR products were restriction digested and mixed with DNA fragments containing a nonpolar Em resistance gene (erm) (24), a nonpolar Km resistance gene (kan), or a polar Ωkan cassette (23) in a ligation reaction to allow for ligation in the following order: the 5′-flanking fragment, followed by the fragment containing the antibiotic resistance gene, and the 3′-flanking fragment. The ligation mixture was introduced into S. sanguinis SK36 by natural transformation (39), and the allelic-exchange event in the antibiotic-resistant transformants was verified by colony PCR with primers located outside the insertion site of the antibiotic resistance gene.

Construction of recombinant S. sanguinis pilB promoter-cat fusion strains.A DNA fragment of 947 bp immediately 5′ to the translation start codon of SSA_2318 was amplified from S. sanguinis SK36 with primers pil_3_BstXI_S and pil_BglII_AS. The PCR product was fused to a promoterless cat gene (40) on pGEM7zf(+), tagged with a Sp resistance gene (spe) (41) at the 5′ end of the fusion, and then established in E. coli DH10B. The resulting plasmid, pTY12, was used as a template to mutate each of putative promoters by inverse PCR. Briefly, primer pairs P1_-10_PstI_S/P1_-10_PstI_AS, P2_-10_PstI_S/P2_-10_PstI_AS, and P3_-10_PstI_S/P3_-10__PstI_AS were used to replace the putative −10 elements of ppilB-1, ppilB-2, and ppilB-3 with a PstI recognition sequence (5′-CTGCAG), respectively. The inverse PCR products were digested, religated, and established in E. coli DH10B. The identity of all recombinant plasmids was confirmed by sequencing analysis. DNA fragments containing the spe-tagged fusions on each of the recombinant plasmids were generated by PCR with primers Spec_BamHI_S and Cat_ClaI_AS and integrated into the chromosome of S. sanguinis SK36 at SSA_1656 using ligation mutagenesis (38). The integration event in the Sp-resistant transformants was confirmed by PCR with primers located outside the insertion site. The resulting strains, ppilB_P1+P2, ppilB_P1+P3, and ppilB_P2+P3, carried fusions with mutated ppilB-3, ppilB-2, and ppilB-1, respectively.

By using the same approach, the plasmid with mutations in one promoter was used as a template to introduce mutations into another promoter. Briefly, primers P1_-10_XhoI_S and P1_-10_XhoI_AS were used to introduce mutations into ppilB-1 in constructs with mutated ppilB-2 and ppilB-3, and primers P2_-10_XhoI_S and P2_-10_XhoI_AS were used to introduce mutations into ppilB-2 in the construct with mutated ppilB-3. The resulting strains, ppilB_P1, ppilB_P2, and ppilB_P3, carried fusions of wild-type ppilB-1, ppilB-2, and ppilB-3, respectively.

CAT assay.Cultures of S. sanguinis strains grown overnight were diluted at 1:20 in prewarmed TH broth and grown to an OD600 of ∼0.6. Cells were harvested, washed once with 10 mM Tris-HCl (pH 7.8), and then resuspended in 1/80 of the culture volume in the same buffer. Cells were homogenized by using a Mini-BeadBeater-8 instrument for a total of 2 min at 4°C. Clear lysates were collected by centrifugation at 10,000 rpm, and the protein concentrations of the lysates were measured using the Bio-Rad protein assay. CAT activity in each lysate was determined according to a method described previously by Shaw (42). The specific activity was calculated as nanomoles of chloramphenicol (Cm) acetylated per minute per milligram of total protein. All reactions were done in triplicate. Reaction mixtures without the addition of Cm were used as a negative control.

Purification of recombinant SSA_2315 protein, production of polyclonal antisera, and Western blot analysis.A DNA fragment containing the sequence encoding amino acids 38 through 151 of SSA_2315 was generated from S. sanguinis SK36 by PCR using the primer pair pil15925BamHIAS/pil15575PstIS and cloned into pQE30 (Qiagen). The cloned fragment was confirmed by sequence analysis. The induction and purification of histidine-tagged SSA_2315 were performed under conditions according to the manufacturer’s instructions (Qiagen). The identity of this recombinant protein was verified by matrix-assisted laser desorption ionization–time of flight mass spectrometry (MALDI-TOF MS). Approximately 2.5 mg of the purified protein was run on a 12% SDS-PAGE gel, and the protein band was excised from the gel and used to generate polyclonal antiserum in rabbits (Genesis). The specificity and titer of the antiserum were assessed by Western blot analysis.

To examine the production of SSA_2315 in S. sanguinis SK36 at different growth phases, cultures at OD600s of 0.3 and 0.6 were harvested and washed once with phosphate-buffered saline (PBS), followed by one wash with 50 mM Tris-HCl (pH 7.0). The cell suspension was resuspended in 1/50 of the culture volume in the same buffer containing 20% sucrose and 2 mM MgCl2. The concentrated cell suspension was treated with 90 U of mutanolysin at 37°C for 1.5 h to release cell wall-anchored proteins, followed by boiling for 25 min to recover all intracellular proteins. One microgram of each sample was loaded onto a 12% SDS-PAGE gel. For Western blot analysis, the separated protein species were transferred to a polyvinylidene difluoride (PVDF) membrane. The membrane was blocked with 10% nonfat milk in PBS containing 0.3% Tween 20 (PBST) overnight at 4°C prior to hybridization with anti-SSA_2315 antibody diluted 1:40,000 in PBST with 5% nonfat dry milk for 1 h. SSA_2315 production was detected with horseradish peroxidase (HRP)-conjugated goat anti-rabbit antibody (GeneTex) and luminol-based Immobilon Western chemiluminescent HRP substrate (Millipore) and imaged on Fuji medical X-ray film.

Immunogold labeling, negative staining, and examination of Tfp by transmission electron microscopy.For immunogold labeling, 3 μl Tris-buffered saline (TBS) (20 mM Tris-HCl [pH 7.4], 150 mM NaCl) was deposited on the top of one freshly grown colony of S. sanguinis SK36 and YC6 on TH agar. The wetted colony was transferred onto a 200-mesh carbon-coated copper grid (Agar) by allowing the grid to make contact with the colony for 30 s. Bacterial cells on the grid were incubated at room temperature with TBS containing 2% bovine serum albumin (BSA) for 1 h, followed by incubation with anti-SSA_2315 antiserum at a 1:50 dilution in TBS for 1 h. At the end of the incubation, the grid was rinsed with drops of TBS, followed by incubation with a goat anti-rabbit IgG conjugate with 10-nm gold particles (Sigma) at room temperature for 45 min. At the end of the incubation, the grid was rinsed gently with drops of TBS and then negatively stained according to a method described previously by Ruiz et al., with 2% phosphotungstic acid at pH 7 (43). The grid was observed using a JEM-1230 electron microscope (JEOL).

Adherence assays.HeLa human cervical cancer cells were seeded in a 24-well plate at 8 × 104 cells per well in Dulbecco’s modified Eagle medium (DMEM) with 10% fetal bovine serum (FBS). SCC-4 human oral squamous cell carcinoma cells were seeded at 7 × 104 cells per well in DMEM–Ham’s F-12 nutrient medium with 10% FBS. Both cell cultures were incubated at 37°C in a 5% CO2 atmosphere until confluence. Exponential-phase cultures of S. sanguinis SK36, YC6, TY10, and TY11 at an OD600 of 0.3 were harvested, resuspended in DMEM or DMEM–Ham’s F-12 medium, and added to the cell monolayer at a multiplicity of infection (MOI) of 100 for 30 min. At the end of the incubation, the nonadherent bacteria were removed, and the monolayers were washed three times with PBS. Two hundred microliters of 0.25% trypsin in 1 mM EDTA was added to each well to detach the cell monolayer. The CFU of attached bacteria were determined by serial dilution and plating. The percentage of adherence was determined as the number of adherent CFU divided by the number of CFU of bacteria added.

Biofilm formation assay.The static biofilm formation of S. sanguinis strains was examined by using a previously described method (44), with modifications. Cultures grown overnight in biofilm medium (BM) (45) containing 40 mM glucose (BMG) were diluted at 1:200 in BMG and transferred into wells of a 96-well polystyrene microtiter plate. The plate was incubated at 37°C in a 10% CO2 atmosphere for 18 h. The biofilm was stained with crystal violet, and the absorbance at 562 nm (OD562) was determined with a microplate reader (SoftMax Pro; Molecular Devices). The final growth yield was a measure of the OD490. All cultures were grown in triplicate.

Statistical analysis.Statistical analysis was performed using analysis of variance (ANOVA) followed by a Dunnett or Tukey test using GraphPad Prism 5. Differences were considered significant if the P value was <0.01.

ACKNOWLEDGMENTS

This work was supported by Ministry of Science and Technology (MOST) of Taiwan grant 105-2320-B-182-015-MY3 to Y.-Y. M. Chen.

We thank R. Faustoferri and S. T. Liu for review of the manuscript.

FOOTNOTES

    • Received 21 November 2018.
    • Accepted 4 January 2019.
    • Accepted manuscript posted online 11 January 2019.

  • Supplemental material for this article may be found at https://doi.org/10.1128/AEM.02788-18.

REFERENCES

  1. 1.↵
    1. Abusleme L,
    2. Dupuy AK,
    3. Dutzan N,
    4. Silva N,
    5. Burleson JA,
    6. Strausbaugh LD,
    7. Gamonal J,
    8. Diaz PI
    . 2013. The subgingival microbiome in health and periodontitis and its relationship with community biomass and inflammation. ISME J 7:1016–1025. doi:10.1038/ismej.2012.174.
    OpenUrlCrossRefPubMedWeb of Science
  2. 2.↵
    1. Gross EL,
    2. Beall CJ,
    3. Kutsch SR,
    4. Firestone ND,
    5. Leys EJ,
    6. Griffen AL
    . 2012. Beyond Streptococcus mutans: dental caries onset linked to multiple species by 16S rRNA community analysis. PLoS One 7:e47722. doi:10.1371/journal.pone.0047722.
    OpenUrlCrossRef
  3. 3.↵
    1. Griffen AL,
    2. Beall CJ,
    3. Campbell JH,
    4. Firestone ND,
    5. Kumar PS,
    6. Yang ZK,
    7. Podar M,
    8. Leys EJ
    . 2012. Distinct and complex bacterial profiles in human periodontitis and health revealed by 16S pyrosequencing. ISME J 6:1176–1185. doi:10.1038/ismej.2011.191.
    OpenUrlCrossRefPubMedWeb of Science
  4. 4.↵
    1. Belda-Ferre P,
    2. Alcaraz LD,
    3. Cabrera-Rubio R,
    4. Romero H,
    5. Simón-Soro A,
    6. Pignatelli M,
    7. Mira A
    . 2012. The oral metagenome in health and disease. ISME J 6:46–56. doi:10.1038/ismej.2011.85.
    OpenUrlCrossRefPubMedWeb of Science
  5. 5.↵
    1. Mylonakis E,
    2. Calderwood SB
    . 2001. Infective endocarditis in adults. N Engl J Med 345:1318–1330. doi:10.1056/NEJMra010082.
    OpenUrlCrossRefPubMedWeb of Science
  6. 6.↵
    1. Forner L,
    2. Larsen T,
    3. Kilian M,
    4. Holmstrup P
    . 2006. Incidence of bacteremia after chewing, tooth brushing and scaling in individuals with periodontal inflammation. J Clin Periodontol 33:401–407. doi:10.1111/j.1600-051X.2006.00924.x.
    OpenUrlCrossRefPubMedWeb of Science
  7. 7.↵
    1. Xu P,
    2. Alves JM,
    3. Kitten T,
    4. Brown A,
    5. Chen Z,
    6. Ozaki LS,
    7. Manque P,
    8. Ge X,
    9. Serrano MG,
    10. Puiu D,
    11. Hendricks S,
    12. Wang Y,
    13. Chaplin MD,
    14. Akan D,
    15. Paik S,
    16. Peterson DL,
    17. Macrina FL,
    18. Buck GA
    . 2007. Genome of the opportunistic pathogen Streptococcus sanguinis. J Bacteriol 189:3166–3175. doi:10.1128/JB.01808-06.
    OpenUrlAbstract/FREE Full Text
  8. 8.↵
    1. Gurung I,
    2. Spielman I,
    3. Davies MR,
    4. Lala R,
    5. Gaustad P,
    6. Biais N,
    7. Pelicic V
    . 1 October 2015. Functional analysis of an unusual type IV pilus in the Gram-positive Streptococcus sanguinis. Mol Microbiol doi:10.1111/mmi.13237.
    OpenUrlCrossRef
  9. 9.↵
    1. Daum B,
    2. Gold V
    . 2018. Twitch or swim: towards the understanding of prokaryotic motion based on the type IV pilus blueprint. Biol Chem 399:799–808. doi:10.1515/hsz-2018-0157.
    OpenUrlCrossRef
  10. 10.↵
    1. Pelicic V
    . 2008. Type IV pili: e pluribus unum? Mol Microbiol 68:827–837. doi:10.1111/j.1365-2958.2008.06197.x.
    OpenUrlCrossRefPubMedWeb of Science
  11. 11.↵
    1. Melville S,
    2. Craig L
    . 2013. Type IV pili in Gram-positive bacteria. Microbiol Mol Biol Rev 77:323–341. doi:10.1128/MMBR.00063-12.
    OpenUrlAbstract/FREE Full Text
  12. 12.↵
    1. Piepenbrink KH,
    2. Sundberg EJ
    . 2016. Motility and adhesion through type IV pili in Gram-positive bacteria. Biochem Soc Trans 44:1659–1666. doi:10.1042/BST20160221.
    OpenUrlAbstract/FREE Full Text
  13. 13.↵
    1. Maldarelli GA,
    2. Piepenbrink KH,
    3. Scott AJ,
    4. Freiberg JA,
    5. Song Y,
    6. Achermann Y,
    7. Ernst RK,
    8. Shirtliff ME,
    9. Sundberg EJ,
    10. Donnenberg MS,
    11. von Rosenvinge EC
    . 1 June 2016. Type IV pili promote early biofilm formation by Clostridium difficile. Pathog Dis doi:10.1093/femspd/ftw061.
    OpenUrlCrossRef
  14. 14.↵
    1. Purcell EB,
    2. McKee RW,
    3. Bordeleau E,
    4. Burrus V,
    5. Tamayo R
    . 2016. Regulation of type IV pili contributes to surface behaviors of historical and epidemic strains of Clostridium difficile. J Bacteriol 198:565–577. doi:10.1128/JB.00816-15.
    OpenUrlAbstract/FREE Full Text
  15. 15.↵
    1. Bordeleau E,
    2. Purcell EB,
    3. Lafontaine DA,
    4. Fortier LC,
    5. Tamayo R,
    6. Burrus V
    . 2015. Cyclic di-GMP riboswitch-regulated type IV pili contribute to aggregation of Clostridium difficile. J Bacteriol 197:819–832. doi:10.1128/JB.02340-14.
    OpenUrlAbstract/FREE Full Text
  16. 16.↵
    1. Varga JJ,
    2. Nguyen V,
    3. O’Brien DK,
    4. Rodgers K,
    5. Walker RA,
    6. Melville SB
    . 2006. Type IV pili-dependent gliding motility in the Gram-positive pathogen Clostridium perfringens and other Clostridia. Mol Microbiol 62:680–694. doi:10.1111/j.1365-2958.2006.05414.x.
    OpenUrlCrossRefPubMedWeb of Science
  17. 17.↵
    1. Varga JJ,
    2. Therit B,
    3. Melville SB
    . 2008. Type IV pili and the CcpA protein are needed for maximal biofilm formation by the Gram-positive anaerobic pathogen Clostridium perfringens. Infect Immun 76:4944–4951. doi:10.1128/IAI.00692-08.
    OpenUrlAbstract/FREE Full Text
  18. 18.↵
    1. Rakotoarivonina H,
    2. Larson MA,
    3. Morrison M,
    4. Girardeau JP,
    5. Gaillard-Martinie B,
    6. Forano E,
    7. Mosoni P
    . 2005. The Ruminococcus albus pilA1-pilA2 locus: expression and putative role of two adjacent pil genes in pilus formation and bacterial adhesion to cellulose. Microbiology 151:1291–1299. doi:10.1099/mic.0.27735-0.
    OpenUrlCrossRefPubMedWeb of Science
  19. 19.↵
    1. Rakotoarivonina H,
    2. Jubelin G,
    3. Hebraud M,
    4. Gaillard-Martinie B,
    5. Forano E,
    6. Mosoni P
    . 2002. Adhesion to cellulose of the Gram-positive bacterium Ruminococcus albus involves type IV pili. Microbiology 148:1871–1880. doi:10.1099/00221287-148-6-1871.
    OpenUrlCrossRefPubMedWeb of Science
  20. 20.↵
    1. Laurenceau R,
    2. Pehau-Arnaudet G,
    3. Baconnais S,
    4. Gault J,
    5. Malosse C,
    6. Dujeancourt A,
    7. Campo N,
    8. Chamot-Rooke J,
    9. Le Cam E,
    10. Claverys JP,
    11. Fronzes R
    . 2013. A type IV pilus mediates DNA binding during natural transformation in Streptococcus pneumoniae. PLoS Pathog 9:e1003473. doi:10.1371/journal.ppat.1003473.
    OpenUrlCrossRefPubMed
  21. 21.↵
    1. Berry JL,
    2. Pelicic V
    . 2015. Exceptionally widespread nanomachines composed of type IV pilins: the prokaryotic Swiss Army knives. FEMS Microbiol Rev 39:134–154. doi:10.1093/femsre/fuu001.
    OpenUrlCrossRefPubMed
  22. 22.↵
    1. Burrows LL
    . 2012. Pseudomonas aeruginosa twitching motility: type IV pili in action. Annu Rev Microbiol 66:493–520. doi:10.1146/annurev-micro-092611-150055.
    OpenUrlCrossRefPubMedWeb of Science
  23. 23.↵
    1. Perez-Casal J,
    2. Caparon MG,
    3. Scott JR
    . 1991. Mry, a trans-acting positive regulator of the M protein gene of Streptococcus pyogenes with similarity to the receptor proteins of two-component regulatory systems. J Bacteriol 173:2617–2624. doi:10.1128/jb.173.8.2617-2624.1991.
    OpenUrlAbstract/FREE Full Text
  24. 24.↵
    1. Rubens CE,
    2. Heggen LM
    . 1988. Tn916 delta E: a Tn916 transposon derivative expressing erythromycin resistance. Plasmid 20:137–142. doi:10.1016/0147-619X(88)90016-9.
    OpenUrlCrossRefPubMedWeb of Science
  25. 25.↵
    1. Sabelnikov AG,
    2. Greenberg B,
    3. Lacks SA
    . 1995. An extended −10 promoter alone directs transcription of the DpnII operon of Streptococcus pneumoniae. J Mol Biol 250:144–155. doi:10.1006/jmbi.1995.0366.
    OpenUrlCrossRefPubMedWeb of Science
  26. 26.↵
    1. Sun Z,
    2. Zhong J,
    3. Liang X,
    4. Liu J,
    5. Chen X,
    6. Huan L
    . 2009. Novel mechanism for nisin resistance via proteolytic degradation of nisin by the nisin resistance protein NSR. Antimicrob Agents Chemother 53:1964–1973. doi:10.1128/AAC.01382-08.
    OpenUrlAbstract/FREE Full Text
  27. 27.↵
    1. Mendez M,
    2. Huang IH,
    3. Ohtani K,
    4. Grau R,
    5. Shimizu T,
    6. Sarker MR
    . 2008. Carbon catabolite repression of type IV pilus-dependent gliding motility in the anaerobic pathogen Clostridium perfringens. J Bacteriol 190:48–60. doi:10.1128/JB.01407-07.
    OpenUrlAbstract/FREE Full Text
  28. 28.↵
    1. Okahashi N,
    2. Nakata M,
    3. Sakurai A,
    4. Terao Y,
    5. Hoshino T,
    6. Yamaguchi M,
    7. Isoda R,
    8. Sumitomo T,
    9. Nakano K,
    10. Kawabata S,
    11. Ooshima T
    . 2010. Pili of oral Streptococcus sanguinis bind to fibronectin and contribute to cell adhesion. Biochem Biophys Res Commun 391:1192–1196. doi:10.1016/j.bbrc.2009.12.029.
    OpenUrlCrossRefPubMed
  29. 29.↵
    1. Giltner CL,
    2. Nguyen Y,
    3. Burrows LL
    . 2012. Type IV pilin proteins: versatile molecular modules. Microbiol Mol Biol Rev 76:740–772. doi:10.1128/MMBR.00035-12.
    OpenUrlAbstract/FREE Full Text
  30. 30.↵
    1. Estrem ST,
    2. Gaal T,
    3. Ross W,
    4. Gourse RL
    . 1998. Identification of an UP element consensus sequence for bacterial promoters. Proc Natl Acad Sci U S A 95:9761–9766.
    OpenUrlAbstract/FREE Full Text
  31. 31.↵
    1. Boekema BK,
    2. Van Putten JP,
    3. Stockhofe-Zurwieden N,
    4. Smith HE
    . 2004. Host cell contact-induced transcription of the type IV fimbria gene cluster of Actinobacillus pleuropneumoniae. Infect Immun 72:691–700. doi:10.1128/IAI.72.2.691-700.2004.
    OpenUrlAbstract/FREE Full Text
  32. 32.↵
    1. Antunes A,
    2. Camiade E,
    3. Monot M,
    4. Courtois E,
    5. Barbut F,
    6. Sernova NV,
    7. Rodionov DA,
    8. Martin-Verstraete I,
    9. Dupuy B
    . 2012. Global transcriptional control by glucose and carbon regulator CcpA in Clostridium difficile. Nucleic Acids Res 40:10701–10718. doi:10.1093/nar/gks864.
    OpenUrlCrossRefPubMedWeb of Science
  33. 33.↵
    1. Richardson AR,
    2. Somerville GA,
    3. Sonenshein AL
    . 2015. Regulating the intersection of metabolism and pathogenesis in Gram-positive bacteria. Microbiol Spectr 3(3):MBP-0004-2014. doi:10.1128/microbiolspec.MBP-0004-2014.
    OpenUrlCrossRef
  34. 34.↵
    1. Willenborg J,
    2. de Greeff A,
    3. Jarek M,
    4. Valentin-Weigand P,
    5. Goethe R
    . 2014. The CcpA regulon of Streptococcus suis reveals novel insights into the regulation of the streptococcal central carbon metabolism by binding of CcpA to two distinct binding motifs. Mol Microbiol 92:61–83. doi:10.1111/mmi.12537.
    OpenUrlCrossRefPubMed
  35. 35.↵
    1. Yang Y,
    2. Zhang L,
    3. Huang H,
    4. Yang C,
    5. Yang S,
    6. Gu Y,
    7. Jiang W
    . 24 January 2017. A flexible binding site architecture provides new insights into CcpA global regulation in Gram-positive bacteria. mBio doi:10.1128/mBio.02004-16.
    OpenUrlAbstract/FREE Full Text
  36. 36.↵
    1. Ota C,
    2. Morisaki H,
    3. Nakata M,
    4. Arimoto T,
    5. Fukamachi H,
    6. Kataoka H,
    7. Masuda Y,
    8. Suzuki N,
    9. Miyazaki T,
    10. Okahashi N,
    11. Kuwata H
    . 20 February 2018. Streptococcus sanguinis noncoding cia-dependent small RNAs negatively regulate expression of type IV pilus retraction ATPase PilT and biofilm formation. Infect Immun doi:10.1128/IAI.00894-17.
    OpenUrlAbstract/FREE Full Text
  37. 37.↵
    1. Chen YY,
    2. Weaver CA,
    3. Mendelsohn DR,
    4. Burne RA
    . 1998. Transcriptional regulation of the Streptococcus salivarius 57.I urease operon. J Bacteriol 180:5769–5775.
    OpenUrlAbstract/FREE Full Text
  38. 38.↵
    1. Lau PC,
    2. Sung CK,
    3. Lee JH,
    4. Morrison DA,
    5. Cvitkovitch DG
    . 2002. PCR ligation mutagenesis in transformable streptococci: application and efficiency. J Microbiol Methods 49:193–205.
    OpenUrlCrossRefPubMedWeb of Science
  39. 39.↵
    1. Paik S,
    2. Senty L,
    3. Das S,
    4. Noe JC,
    5. Munro CL,
    6. Kitten T
    . 2005. Identification of virulence determinants for endocarditis in Streptococcus sanguinis by signature-tagged mutagenesis. Infect Immun 73:6064–6074. doi:10.1128/IAI.73.9.6064-6074.2005.
    OpenUrlAbstract/FREE Full Text
  40. 40.↵
    1. Horinouchi S,
    2. Weisblum B
    . 1982. Nucleotide sequence and functional map of pC194, a plasmid that specifies inducible chloramphenicol resistance. J Bacteriol 150:815–825.
    OpenUrlAbstract/FREE Full Text
  41. 41.↵
    1. LeBlanc DJ,
    2. Lee LN,
    3. Inamine JM
    . 1991. Cloning and nucleotide base sequence analysis of a spectinomycin adenyltransferase AAD(9) determinant from Enterococcus faecalis. Antimicrob Agents Chemother 35:1804–1810.
    OpenUrlAbstract/FREE Full Text
  42. 42.↵
    1. Shaw WV
    . 1975. Chloramphenicol acetyltransferase from chloramphenicol-resistant bacteria. Methods Enzymol 43:737–755.
    OpenUrlCrossRefPubMed
  43. 43.↵
    1. Ruiz T,
    2. Lenox C,
    3. Radermacher M,
    4. Mintz KP
    . 2006. Novel surface structures are associated with the adhesion of Actinobacillus actinomycetemcomitans to collagen. Infect Immun 74:6163–6170. doi:10.1128/IAI.00857-06.
    OpenUrlAbstract/FREE Full Text
  44. 44.↵
    1. O’Toole GA,
    2. Kolter R
    . 1998. Initiation of biofilm formation in Pseudomonas fluorescens WCS365 proceeds via multiple, convergent signalling pathways: a genetic analysis. Mol Microbiol 28:449–461. doi:10.1046/j.1365-2958.1998.00797.x.
    OpenUrlCrossRefPubMedWeb of Science
  45. 45.↵
    1. Loo CY,
    2. Corliss DA,
    3. Ganeshkumar N
    . 2000. Streptococcus gordonii biofilm formation: identification of genes that code for biofilm phenotypes. J Bacteriol 182:1374–1382. doi:10.1128/JB.182.5.1374-1382.2000.
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KEYWORDS

CcpA
Streptococcus sanguinis
adherence
biofilms
type IV pilus

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