Host Response and Inflammation

Enterococcus faecalis Demonstrates Pathogenicity through Increased Attachment in an Ex Vivo Polymicrobial Pulpal Infection

Wayne Nishio Ayre, Genevieve Melling, Camille Cuveillier, Madhan Natarajan, Jessica L. Roberts, Lucy L. Marsh, Christopher D. Lynch, Jean-Yves Maillard, Stephen P. Denyer, Alastair J. Sloan
Nancy E. Freitag, Editor
DOI: 10.1128/IAI.00871-17

ABSTRACT

This study investigated the host response to a polymicrobial pulpal infection consisting of Streptococcus anginosus and Enterococcus faecalis, bacteria commonly implicated in dental abscesses and endodontic failure, using a validated ex vivo rat tooth model. Tooth slices were inoculated with planktonic cultures of S. anginosus or E. faecalis alone or in coculture at S. anginosus/E. faecalis ratios of 50:50 and 90:10. Attachment was semiquantified by measuring the area covered by fluorescently labeled bacteria. Host response was established by viable histological cell counts, and inflammatory response was measured using reverse transcription-quantitative PCR (RT-qPCR) and immunohistochemistry. A significant reduction in cell viability was observed for single and polymicrobial infections, with no significant differences between infection types (∼2,000 cells/mm2 for infected pulps compared to ∼4,000 cells/mm2 for uninfected pulps). E. faecalis demonstrated significantly higher levels of attachment (6.5%) than S. anginosus alone (2.3%) and mixed-species infections (3.4% for 50:50 and 2.3% for 90:10), with a remarkable affinity for the pulpal vasculature. Infections with E. faecalis demonstrated the greatest increase in tumor necrosis factor alpha (TNF-α) (47.1-fold for E. faecalis, 14.6-fold for S. anginosus, 60.1-fold for 50:50, and 25.0-fold for 90:10) and interleukin 1β (IL-1β) expression (54.8-fold for E. faecalis, 8.8-fold for S. anginosus, 54.5-fold for 50:50, and 39.9-fold for 90:10) compared to uninfected samples. Immunohistochemistry confirmed this, with the majority of inflammation localized to the pulpal vasculature and odontoblast regions. Interestingly, E. faecalis supernatant and heat-killed E. faecalis treatments were unable to induce the same inflammatory response, suggesting E. faecalis pathogenicity in pulpitis is linked to its greater ability to attach to the pulpal vasculature.

INTRODUCTION

The dental pulp is a complex environment composed of soft connective tissue, nerves, blood vessels, and a variety of cells, such as dental pulp stem cells, fibroblasts, and odontoblasts (1). When the pulp becomes inflamed in response to bacterial infection or other stimuli, this is known as pulpitis. Early stages are considered “reversible,” and treatment involves removal of the stimulus, such as carious lesions, in order to maintain pulp vitality. If untreated, however, the microbial invasion may progress into the deeper dentine and subsequently the pulpal chamber, resulting in severe tissue degradation and necrosis. This condition, known as “irreversible pulpitis,” requires a challenging and difficult endodontic or root canal treatment, which involves the removal of the pulp and obturation with an inert material. The success rate of root canal treatments is highly variable, ranging from 31% to 96% depending on clinical considerations (2), and studies across a range of countries have shown that a high percentage (up to 67.9%) of patients who have undergone this treatment subsequently develop apical periodontitis (3, 4). An alternative endodontic treatment is vital pulpotomy, which involves removal of the coronal pulp, leaving the radicular pulp vital and free of any pathological alterations (5). Although this procedure is thought to require shorter appointment times and can be accomplished in one visit, the efficacy of the technique is debated, with success rates of clinical studies ranging from 70% to 96% (6). Accurate models to better understand the process of pulpal infection and to test the efficacy of novel therapeutics will aid in the development of more effective vital pulp treatments. In vitro monolayer cell culture models lack the complexity of the pulpal matrix, while in vivo studies suffer from systemic factors, high costs, and ethical considerations. To overcome these limitations, Roberts et al. (7) developed an ex vivo coculture system to model pulpal infections on rat tooth slices. This study focused predominantly on the Streptococcus anginosus group (SAG), consisting of S. anginosus, Streptococcus constellatus and Streptococcus intermedius, Gram-positive cocci that are part of the body's commensal flora. The members of this group are known to be primary colonizers of the oral cavity due to their ability to attach to the salivary pellicle and other oral bacteria (8). They are considered opportunistic pathogens and have been reported to form dental abscesses (9). The study by Roberts et al. demonstrated a significant reduction in viable pulp cells and an increase in cytokine expression and bacterial attachment over 24 h as a result of S. anginosus infections (7).

Although Roberts et al. demonstrated invasion of the dental pulp by S. anginosus group species, the number of microbial species encountered in the oral cavity is far more diverse, with studies identifying between 100 and 300 different species from different regions of the oral cavity in healthy individuals (10). It is therefore unsurprising that complex mixed-species microbiomes are often detected in cases of pulpitis (11). As lesions progress into the tooth, a shift in microbial species due to environmental and nutritional changes has been well documented (12). Of particular interest is the species Enterococcus faecalis, a Gram-positive facultative anaerobic coccus that is also part of the normal human commensal flora (13). E. faecalis has been shown to be pathogenic, particularly in endodontic failure (14), with prevalence in such infections ranging from 24% up to 77% (15). Although highly implicated in persistent endodontic failure, molecular studies have recently revealed that the species is frequently present in necrotic pulps, highlighting its potential role in late-stage pulpitis (16, 17).

This study aimed to use a validated ex vivo coculture model to quantify and better understand the host tissue response to mixed-species pulpal infections caused by S. anginosus and E. faecalis. Understanding the mechanism of complex pulpal infections and the host inflammatory response may elucidate potential targets for more effective vital pulp therapies.

RESULTS

Mixed-species culture does not significantly influence S. anginosus and E. faecalis growth rates.The growth characteristics of a simple mixed-species planktonic broth culture were investigated to ensure potential competitive growth between S. anginosus and E. faecalis would not influence the ex vivo experiments investigating host tissue response.

Clinical isolates of the species S. anginosus and E. faecalis were selected from the culture collection of the Oral Microbiology Unit, School of Dentistry at Cardiff University. Species identities were confirmed by standard microbial identification tests and 16S rRNA sequencing, as described in Materials and Methods and the supplemental material (see Fig. S1 and S2 in the supplemental material).

Figure 1 shows the planktonic growth curves for S. anginosus and E. faecalis alone and in combination at ratios of 50:50 and 90:10 over 24 h in brain heart infusion (BHI) broth. E. faecalis reached mid-log phase earlier than S. anginosus (8 h for E. faecalis compared to 10 h for S. anginosus). When cultured at a ratio of 50:50, however, S. anginosus reached mid-log phase at a time similar to that for E. faecalis (10 h). When the bacteria were cultured at an S. anginosus/E. faecalis ratio of 90:10, S. anginosus reached mid-log phase at approximately 8 h and E. faecalis at approximately 12 h. Growth rate calculations during the log phase demonstrated no significant differences between E. faecalis and S. anginosus under all culture conditions (P > 0.05) (Table 1).

FIG 1
FIG 1

Growth curves of E. faecalis (A), S. anginosus (B), and E. faecalis and S. anginosus combined at ratios of 50:50 (C) and 90:10 (D). Mean values of three experimental repeats are shown, with error bars indicating standard deviations.

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TABLE 1

Growth rates of S. anginosus and E. faecalis during log phase, alone and in combination at ratios of 50:50 and 90:10

E. faecalis demonstrates greater levels of attachment to dental pulp than S. anginosus at 24 h, with particular affinity for the pulpal vasculature.To assess differences in bacterial attachment to the dental pulp, the ex vivo rat tooth model was infected with planktonic cultures of S. anginosus and E. faecalis individually or as mixed-species infections. Gram staining and fluorescent labeling of bacteria were undertaken to localize and semiquantify bacterial attachment.

High levels of bacterial attachment to the pulp were detected for tooth slices incubated with E. faecalis (Fig. 2A) and mixed species (S. anginosus and E. faecalis) (Fig. 2B and C). Attachment was predominantly observed in intercellular spaces within the pulpal matrix and around the pulpal vasculature. Bacteria were also observed attached to soft tissue surrounding the tooth and within dentinal tubules (Fig. 2D and E). Attachment of bacteria was not detected using Gram staining on tooth slices incubated with S. anginosus alone.

FIG 2
FIG 2

Gram stains of tooth slices infected with E. faecalis (A), 50:50 S. anginosus-E. faecalis (B), and 90:10 S. anginosus-E. faecalis (C to E). The arrows indicate areas of bacterial attachment. P, dental pulp; D, dentine; S, soft tissue surrounding the tooth. Representative images of three experimental repeats are shown.

Control samples demonstrated low levels of background fluorescence (Fig. 3A). Infections consisting of E. faecalis alone had the greatest fluorescent signal, in particular centered near the pulpal vasculature (Fig. 3B). S. anginosus demonstrated low bacterial attachment spread evenly across the pulp (Fig. 3C). When, E. faecalis and S. anginosus were combined, higher levels of attachment were observed than with S. anginosus alone (Fig. 3D and E), with attachment again localized predominantly to the pulpal vasculature. When the percent bacterial coverage was semiquantified (Fig. 3F), the single-species E. faecalis infection had significantly higher levels of bacterial attachment than S. anginosus alone (approximately 6.5% compared to 2%; P = 0.00021) and the mixed-species infections (50:50, P = 0.0235; 90:10, P = 0.0032).

FIG 3
FIG 3

(A to E) Localization of bacterial attachment by fluorescence microscopy for tooth slices infected with no-bacteria control (A), E. faecalis (B), S. anginosus (C), 50:50 S. anginosus-E. faecalis (D), and 90:10 S. anginosus-E. faecalis (E). P, dental pulp; O, odontoblast region; D, dentine. Representative images of three experimental repeats are shown. (F) Bacterial coverage as quantified by the area of fluorescence relative to the total pulp area (*, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001). Mean values of three experimental repeats are shown, with error bars indicating standard errors of the mean.

S. anginosus and E. faecalis infections significantly reduce pulp cell viability, with E. faecalis infections inducing a significantly greater inflammatory response.To establish the dental pulp host responses to S. anginosus and E. faecalis infections alone and as mixed-species infections, histomorphometric analysis was performed alongside reverse transcription-quantitative PCR (RT-qPCR) and immunohistochemistry for tumor necrosis factor alpha (TNF-α) and interleukin 1β (IL-1β) expression.

Histological cell counts of the infected tooth sections demonstrated a significant reduction (P ≤ 0.05) in viable cells due to infection by both E. faecalis and S. anginosus alone and in combination (Fig. 4A). There were no significant differences in cell numbers between single-species infections and multispecies infections.

FIG 4
FIG 4

(A) Viable cells counted per square millimeter of pulp. Tooth slices infected with E. faecalis and S. anginosus, both alone and in combination, after 24 h all resulted in a significant reduction in viable-cell numbers in the pulp compared to the noninfected control (*, P ≤ 0.05). Mean values of three experimental repeats are shown, with error bars indicating standard errors of the mean. (B and C) Fold change in TNF-α (B) and IL-1β (C) gene expression as a result of E. faecalis and S. anginosus infections, alone and in combination (*, P ≤ 0.05, and **, P ≤ 0.01 compared to control samples; +, P ≤ 0.05, and ++, P ≤ 0.01). Mean values of three experimental repeats are shown, with error bars indicating standard errors of the mean. (D) Immunohistochemistry of TNF-α and IL-1β for control samples and tooth slices infected with S. anginosus, E. faecalis, 50:50 S. anginosus-E. faecalis, and 90:10 S. anginosus-E. faecalis. Representative images of three experimental repeats are shown.

All infected samples had significantly higher proinflammatory cytokine expression (TNF-α [Fig. 4B] and IL-1β [Fig. 4C]) than the control samples (P ≤ 0.05). The single-species infection with E. faecalis resulted in significantly higher levels of TNF-α and IL-1β expression than S. anginosus infection (P = 0.0276 and P = 0.0234 for TNF-α and IL-1β, respectively). Combining E. faecalis and S. anginosus did not result in a significantly higher inflammatory response from the pulp than E. faecalis alone (TNF-α, P = 0.493 and P = 0.096 for 50:50 and 90:10, respectively; IL-1β, P = 0.988 and P = 0.400 for 50:50 and 90:10, respectively).

Negative controls replacing the primary TNF-α antibody with a nonimmune immunoglobulin G (IgG) control showed no immunopositivity (see Fig. S3 in the supplemental material). Similarly, primary exclusion controls were negative for staining, indicating specific binding of the secondary antibody (see Fig. S3 in the supplemental material). Control samples demonstrated low expression of TNF-α, and interestingly, S. anginosus alone did not induce a high TNF-α response (Fig. 4D). Samples incubated with E. faecalis alone or in combination with S. anginosus had the most pronounced staining within both the pulp (around the vasculature) and the odontoblast layer. The levels of TNF-α staining in these samples were similar to those encountered in the rat lung positive control (see Fig. S3 in the supplemental material).

Immunohistochemistry staining for IL-1β showed no positive signal for IgG and the primary exclusion controls (see Fig. S3 in the supplemental material). Similar to the TNF-α immunohistochemistry, the control sample and the sample incubated with S. anginosus alone had few positively stained cells, while samples incubated with E. faecalis alone and in combination with S. anginosus had more positively stained cells (Fig. 4D). Although the level of staining was not as pronounced as that observed with TNF-α, the positive cells were again located adjacent to the pulpal vasculature and similar in staining to the positive lung control (see Fig. S3 in the supplemental material).

Greater host inflammatory response to E. faecalis is not due to differences in water-soluble cell wall proteins or culture supernatants.To establish whether the increased host inflammatory response to E. faecalis was due to specific water-soluble cell proteins or components of the culture supernatant, SDS-PAGE was performed to identify proteins in water-soluble cell wall proteins and culture supernatants. Similarly, heat-killed E. faecalis and the E. faecalis supernatant were used to stimulate the pulp in order to assess the host response.

Few differences were observed between the water-soluble cell wall proteins of S. anginosus and E. faecalis when cultured alone and in combination with each other (see Fig. S4A in the supplemental material). In terms of the culture supernatant, there was one band at approximately 35 kDa observed with the E. faecalis cultures that was not observed with S. anginosus (see Fig. S4B in the supplemental material).

When culturing the rat tooth slices with the E. faecalis supernatant or heat-killed E. faecalis, no significant differences were observed in TNF-α expression compared to the untreated controls (Fig. 5A) (P = 0.196 and P = 0.152 for supernatant and heat-killed E. faecalis, respectively). A significant increase was observed in IL-1β expression for the tooth slices cultured with heat-killed E. faecalis compared to the untreated controls (Fig. 5B) (P = 0.041), but not for E. faecalis supernatant (P = 0.148).

FIG 5
FIG 5

(A and B) Fold change in TNF-α (A) and IL-1β (B) gene expression relative to β–actin as a result of treating tooth slices with E. faecalis supernatant and heat-killed E. faecalis (*, P ≤ 0.05 compared to control samples). Mean values of three experimental repeats are shown, with error bars indicating standard errors of the mean. (C) Immunohistochemistry of TNF-α and IL-1β for control samples and tooth slices infected with E. faecalis supernatant and heat-killed E. faecalis. Representative images of three experimental repeats are shown.

The negative controls (IgG and primary exclusion controls) and the control sample for the TNF-α immunohistochemistry did not show staining (see Fig. S5 in the supplemental material). The tooth slices incubated with E. faecalis supernatant had few cells stained positive for TNF-α, the majority of which were concentrated at the pulpal vasculature and the odontoblast layer (Fig. 5C). Similarly, heat-killed E. faecalis had few cells expressing TNF-α (Fig. 5C), while the lung positive control stained positive for TNF-α (see Fig. S5 in the supplemental material).

The IgG control, the primary exclusion control, and the untreated sample (see Fig. S5 in the supplemental material) did not stain positive for IL-1β. Fewer cells were positive for IL-1β than for TNF-α (Fig. 5C). Samples treated with E. faecalis supernatant showed some cells stained positive within the pulpal vasculature, while heat-killed E. faecalis showed few positively stained cells. The positive lung control demonstrated cells stained positive for IL-1β expression (see Fig. S5 in the supplemental material).

DISCUSSION

This study successfully employed an existing ex vivo rat tooth infection model to study the effect of mixed-species E. faecalis and S. anginosus pulpal infections on cell viability, bacterial attachment, and host inflammatory response.

By studying simple planktonic growth kinetics, it was established that E. faecalis caused the S. anginosus bacteria to reach log phase at a higher rate. This concept of polymicrobial synergy has been highlighted in recent work, which investigated metabolite cross-feeding, where metabolic end products produced by one bacterium are consumed by a second community member (18–20). In particular, this has been demonstrated for a similar oral pathogen, Streptococcus gordonii. Lactate produced by S. gordonii as the primary metabolite during catabolism of carbohydrates was found to support the growth of Aggregatibacter actinomycetemcomitans (20). Interestingly, in a study using a primate model, the addition of E. faecalis to a four-strain mixed-species culture resulted in higher levels of survival of all four strains of bacteria than in the absence of E. faecalis (21). Another mechanism for coordinating activities and communicating between microbial species is quorum sensing, which has been shown to occur between different groups of streptococci (22). Although the rate of growth during the log phase was not altered during mixed-species planktonic culture in this study, it is important to appreciate that under mixed-species biofilm conditions, alterations in growth are likely to occur.

The mixed-species infection did not result in higher levels of bacterial attachment than with E. faecalis alone. The data suggest that E. faecalis is capable of attaching to the dental pulp to a greater extent than S. anginosus, with a particular affinity for the pulpal vasculature. This was not attributed to a higher rate of growth or a higher number of bacteria, as a similar number of S. anginosus cells were counted after 24 h in planktonic broth culture. Similarly, in the mixed-species culture, where S. anginosus achieved log phase at an earlier time point, attachment was not as high as for E. faecalis alone. The increased attachment may therefore be due to differences between the species in terms of motility, sensing, or cell surface adhesins. E. faecalis and S. anginosus are classified as groups D and F, respectively, using Lancefield grouping (23), a method of grouping based on the carbohydrate antigens on the cell wall. These differences in surface carbohydrates could mediate changes in attachment to epithelial cells, as demonstrated by Guzmàn et al. (24). A review by Fisher and Phillips (25) highlighted E. faecalis-specific cell wall components that play a vital role in pathogenic adhesion. Aggregation substance (Agg) increases hydrophobicity and aids adhesion to eukaryotic and prokaryotic surfaces and also encourages the formation of mixed-species biofilms through adherence to other bacteria. Extracellular surface protein (ESP) promotes adhesion, antibiotic resistance, and biofilm formation. Adhesin to collagen of E. faecalis (ACE) is a collagen binding protein belonging to the microbial surface components recognizing adhesive matrix molecules (MSCRAMM) family. ACE plays a role in the pathogenesis of endocarditis, and E. faecalis mutants that do not express ACE have been shown to have significantly reduced attachment to collagen types I and IV but not fibrinogen (26, 27). While S. anginosus has been shown to adhere to the extracellular matrix components fibronectin, fibrinogen, and laminin, binding to collagen types I and IV was much less prominent (28). This is of particular interest in explaining differences in pulpal adherence and the tendency of E. faecalis to localize near the pulpal vasculature, as collagen fibers are often found at higher density around blood vessels and nerves (29).

Although the levels of cell death were the same in the groups tested, infections consisting of E. faecalis alone produced a greater inflammatory response than S. anginosus and mixed-species infections. This increase in inflammation was not due to supernatant or water-soluble cell wall virulence factors of E. faecalis, as treatment of the dental pulp with these isolated factors did not yield high levels of TNF-α and IL-1β expression at gene and protein levels. Basic analysis of supernatant and water-soluble cell wall proteins by SDS-PAGE showed similar bands; however, this might have been due to the absence of serum or collagen (present in the coculture model), which have been shown to influence the production of virulence factors, such as ACE (27). These results indicate the pulpal inflammation caused by E. faecalis is likely due to the higher levels of attachment to the dental pulp. Similar pathogenic traits have been established for E. faecalis in urinary tract infections and endocarditis (30). Increased attachment to the dental pulp would allow direct contact between cells and cell wall components, such as lipoteichoic acid (LTA), which induces activation of cluster of differentiation 14 (CD-14) and Toll-like receptor 2 (TLR-2) (31). An in vivo study that infected canine pulp with lipopolysaccharides (LPS) from Escherichia coli and LTA from E. faecalis demonstrated that LTA treatment led to pulp destruction, albeit to a lesser extent than LPA (32). In vitro studies investigating macrophage responses to E. faecalis LTA found that TNF-α expression was significantly increased in a dose-dependent manner (33), with one study attributing it to the NF-κB and p38 MAPK signaling pathways (34). These studies, however, were performed using monolayer cultures, allowing easy access for LTA to activate Toll-like receptors, whereas the presence of extracellular matrix would limit the penetration of virulence factors into the dental pulp in vivo. Furthermore, macrophages are normally present as monocytes in normal healthy pulp and require a stimulus to become activated (35). Studies using immunohistochemistry have shown these monocytes, as well as dendritic cells, to be located predominantly around blood vessels, with few distributed throughout the pulp (36, 37).

High levels of TNF-α expression were also observed in the odontoblast region using immunohistochemistry. Due to their anatomical location, odontoblasts are the first cells to encounter foreign antigens through either infiltration of virulence factors through dentinal tubules or the breakdown of enamel and dentine. By Gram staining in this study, E. faecalis was observed within the dentinal tubules of the infected tooth slices. This phenomenon has been previously reported in human teeth (38). Odontoblasts, which line the dentine, have been shown to express TLRs and to play a role in the pulp's immune response, in particular to bacterial exotoxins (39–41). This explains the high inflammatory response observed for both infections and supernatant treatments when assessed using immunohistochemistry. Cytokine gene expression using RT-qPCR, however, did not demonstrate higher levels when the dental pulp was treated with supernatants or heat-killed bacteria. This may be attributed to the fact that the methods employed for pulp extraction would be unlikely to fully remove the odontoblast cells.

Although the host response to a mixed-species infection consisting of S. anginosus and E. faecalis was established and the potential pathogenicity of E. faecalis in pulpal infections was elucidated, there are several limitations to this study. The methods employed to fluorescently localize the bacteria could potentially result in diffusion-related artifacts. More specific postprocessing techniques, such as fluorescent in situ hybridization (FISH) probes, may allow more specific identification, quantification, and localization of mixed-species pulpal infections. While the ex vivo model offers a three-dimensional (3D) organotypic culture setting, the static nature, which lacks blood flow, does not allow full observation of the systemic immune response. Potential methods to overcome this may involve addition of monocytes directly to the culture media and prolonged incubation times to stimulate repair mechanisms. Closer examination of attachment mechanisms using ACE-negative E. faecalis mutants and purified LTA would also help fully establish the pathogenicity of E. faecalis in pulpal infections. This will allow the model to be used to develop more effective treatments for pulpitis by assessing the efficacy of antimicrobial and anti-inflammatory treatments in inhibiting bacterial colonization.

In conclusion, this study modeled a mixed-species pulpal infection consisting of S. anginosus and E. faecalis using a validated ex vivo rat tooth model. Although E. faecalis caused S. anginosus to reach the logarithmic growth phase more rapidly, the mixed-species infection did not result in higher levels of cell death, attachment, or inflammatory response from the dental pulp. E. faecalis was found to elicit a much greater inflammatory response, which was due to higher levels of attachment to the dental pulp, with a particular affinity for the pulpal vasculature. Future work will focus on assessing the mechanisms and attachment kinetics in order to elucidate the molecular process and the rate at which E. faecalis colonizes the pulp.

MATERIALS AND METHODS

Materials.All reagents, including culture media, broths, and agars, were purchased from Thermo Scientific (Leicestershire, UK) unless otherwise stated.

Bacterial identification.The S. anginosus and E. faecalis strains studied were clinical isolates selected from the culture collection of the Oral Microbiology Unit, School of Dentistry at Cardiff University. To confirm the identity of the species, standard microbial identification tests were performed by assessing the colony appearance on blood agar, Gram staining, hemolysis, presence of catalase, lactose fermentation (MacConkey agar), Lancefield grouping, and bile esculin agar growth.

16S rRNA sequencing was also performed on the S. anginosus and E. faecalis clinical isolates to validate species identity. S. anginosus and E. faecalis were cultured overnight in BHI broth at 37°C and 5% CO2. DNA was extracted using a QIAamp DNA minikit (Qiagen, Manchester, UK), according to the manufacturer's instructions. The DNA was used in a PCR with 16S rRNA bacterial universal primers D88 (F primer; 5′-GAGAGTTTGATYMTGGCTCAG-3′) and E94 (R primer; 5′-GAAGGAGGTGWTCCARCCGCA-3′) (42), and sequencing of the products was performed by Central Biotechnology Services (Cardiff University) using a 3130xl genetic analyzer (Applied Biosystems). DNA sequences were aligned with GenBank sequences using BLAST (NCBI) to establish the percent sequence identity.

Growth curves.Overnight cultures of S. anginosus and E. faecalis in BHI broth were prepared and diluted to 108 CFU/ml (absorbance at 600 nm, 0.08 to 0.1). The inoculum was diluted in BHI broth to give a starting concentration of 102 CFU/ml. Mixed-species planktonic cultures with a total of 102 CFU/ml consisting of 50% S. anginosus and 50% E. faecalis (referred to here as 50:50) and 90% S. anginosus and 10% E. faecalis (referred to here as 90:10) were prepared. The broths were incubated at 37°C and 5% CO2, and 1-ml aliquots were removed every 4 h for 24 h. The absorbance of the aliquots was measured at 600 nm using an Implen (Munich, Germany) OD600 DiluPhotometer, and 50 μl was spiral plated on tryptic soya agar using a Don Whitley (West Yorkshire, UK) automated spiral plater. The remaining aliquot was then heat treated at 60°C for 30 min prior to spiral plating on bile esculin agar containing 6.5% (wt/wt) sodium chloride. Heat treatment and the presence of high concentrations of bile and sodium chloride permit the growth of only E. faecalis and not S. anginosus (43). The plates were incubated at 37°C and 5% CO2 for 24 h prior to counting. The E. faecalis counts were subtracted from the total counts to give the number of S. anginosus bacteria. The specific growth rate was calculated using the log phase of each growth curve and the following equation: μ = ln(x − x0)/t, where μ is the growth rate in CFU per milliliter per hour, x is the number of CFU per milliliter at the end of the log phase, x0 is the number of CFU per milliliter at the start of the log phase, and t is the duration of the log phase in hours.

Coculture model.The coculture rat tooth infection model was prepared as described by Roberts et al. (7). Twenty-eight-day-old male Wistar rats were sacrificed under schedule 1 of the UK Animals Scientific Procedures Act, 1986, by a qualified technician at the Joint Biological Services Unit, Cardiff University, for harvesting of tissue. Upper and lower incisors were extracted, and the incisors were cut into 2-mm-thick transverse sections using a diamond-edged rotary bone saw (TAAB, Berkshire, UK). The sections were transferred to fresh sterile Dulbecco's modified Eagle medium (DMEM) for no more than 20 min before being cultured in 2 ml DMEM supplemented with 10% (vol/vol) heat-inactivated fetal calf serum, 0.15 mg/ml vitamin C, 200 mmol/liter l-glutamine, 100 U/ml penicillin, 100 μg/ml streptomycin sulfate, and 250 ng/ml amphotericin B at 37°C and 5% CO2 for 24 h. The tooth slices were then washed in 2 ml of phosphate-buffered saline (PBS), transferred to supplemented DMEM without antibiotics, and incubated overnight to remove traces of antibiotic. S. anginosus and E. faecalis were cultured to log phase in BHI broth for 8 to 12 h before dilution to 102 CFU/ml in BHI broth. The bacteria were then used alone or combined for mixed-species infections (S. anginosus/E. faecalis ratios of 50:50 and 90:10). Forty microliters of 1% (wt/vol) fluorescein diacetate (FDA) in acetone was added to 2 ml of the bacterial suspension and incubated for 30 min at 37°C and 5% CO2 before being passed through a 0.22-μm syringe-driven filter unit (Millipore, Oxford, UK). Bacteria captured on the filter were then resuspended in 2 ml sterile supplemented DMEM without antibiotics and with 10% (vol/vol) BHI (referred to here as DMEM-BHI) and used to inoculate one tooth slice. The tooth slices were incubated with the bacteria at 37°C and 5% CO2 for 24 h under constant agitation at 60 rpm in the dark. Sterile DMEM-BHI was used as a control. After incubation, the tooth slices were processed for histology in the dark. The tooth slices were fixed in 10% (wt/vol) neutral buffered formalin at room temperature for 24 h. The slices were demineralized in 10% (wt/vol) formic acid at room temperature for 72 h; dehydrated through a series of 50% (vol/vol), 70% (vol/vol), 95% (vol/vol), and 100% (vol/vol) ethanol, followed by 100% (vol/vol) xylene, for 5 min each; and embedded in paraffin wax. Sections 5 μm thick were cut and viewed under a fluorescence microscope with a fluorescein isothiocyanate (FITC) filter, with images captured using a Nikon digital camera and ACT-1 imaging software (Nikon UK Ltd., Surrey, UK). To quantify cell viability and structural degradation, sections were stained with hematoxylin and eosin (H&E) prior to capturing images with a light microscope.

Gram stain of tissue sections.Gram stains of tooth slices were performed using a modified Brown and Brenn method (44). Paraffin-embedded tooth slices were cut, using a microtome, into 5-μm sections and rehydrated through a series of xylene and 100, 95, and 70% (vol/vol) ethanol for 5 min each. The sections were immersed in 0.2% (wt/vol) crystal violet for 1 min, rinsed with distilled water, immersed in Gram's iodine for 1 min, rinsed with distilled water, decolorized with acetone for 5 s, and counterstained for 1 min with basic fuchsin solution prior to washing with distilled water and mounting. Light microscopy images were captured at ×100 magnification using a Nikon digital camera and ACT-1 imaging software (Nikon UK Ltd., Surrey, UK).

Semiquantification of cell viability by cell counts.ImageJ (National Institutes of Health, Bethesda, MD, USA) was used to count the nuclei per pulp on stained histological sections. For each time point, sections were cut from 5 tooth slices. Images were captured at ×20 magnification and combined using ImageJ software (see Fig. S6 in the supplemental material). The blue field was extracted from the images, and the moments threshold method was applied to separate the pulp cells. The watershed function was applied to split adjacent cell nuclei, and the particles ranging in size from 3 to 100 μm2 were counted. The data were normalized to the pulpal area, and standard errors of the mean were calculated.

Semiquantification of bacterial coverage.ImageJ was used to quantify the area of the pulp inoculated with fluorescent bacteria. The green field of the fluorescent image was extracted, and the image was converted into binary form using the moments threshold method. The pulpal area was manually selected, and the total area of the pulp was measured. The area covered by fluorescent bacteria was then measured and calculated as a percentage of the selected pulp area (see Fig. S7 in the supplemental material).

RT-qPCR of cytokines.Four-millimeter-thick tooth slices were cultured as previously described for 24 h with either sterile DMEM-BHI as a control, DMEM-BHI inoculated with 102 CFU/ml S. anginosus or E. faecalis, or DMEM-BHI with mixed species (S. anginosus and E. faecalis at 50:50 and 90:10 ratios). After incubation, the tooth slices were transferred to sterile PBS, and the pulp was removed by flushing the pulpal cavity with PBS using a 0.1-mm needle and syringe. RNA was extracted using TRIzol reagent (ThermoFisher Scientific, Loughborough, UK), followed by RNase treatment (Promega, Southampton, UK) according to the manufacturers' instructions.

Analysis of gene expression was performed in accordance with the Minimum Information for Publication of Quantitative Real-Time PCR Experiments (MIQE) guidelines (45). RNA concentrations were determined using a NanoVue spectrophotometer (GE Healthcare Life Sciences, Buckinghamshire, UK). RNA purity was determined by ensuring the ratio of absorbance at 260/280 nm was above 1.8, and RNA quality was checked by separating 1 μg of RNA electrophoretically on a 2% agarose gel containing SafeView (NBS Biologicals, Cambridgeshire, UK) in Tris-borate-EDTA buffer to ensure intact 28S and 18S rRNA bands using a Gel Doc EZ system (Bio-Rad, Hertfordshire, UK). Figure S8 in the supplemental material shows RNA integrity following extraction for the samples tested.

cDNA was synthesized by reverse transcription using Promega (Southampton, UK) reagents in a G-Storm (Somerton, UK) GS1 thermocycler. One microgram extracted RNA was combined with 1 μl random primer in a 15-μl reaction mixture in nuclease-free water at 70°C for 5 min. The suspension was added to 5 μl Moloney murine leukemia virus (MMLV) reaction buffer (1.25 μl deoxyribonucleotide triphosphates [dNTPs] [10 mM stock dNTPs], 0.6 μl RNasin, 1 μl MMLV enzyme, and 2.15 μl nuclease-free water) and incubated at 37°C for 1 h.

The resultant cDNA was diluted 1:10 in nuclease-free water (25 ng cDNA). The forward and reverse primers used are listed in Table 2. Ten microliters of PrecisionFast qPCR SYBR green MasterMix with low ROX (PrimerDesign, Chandler's Ford, United Kingdom) was combined with 2 μl of forward and 2 μl of reverse primers (3 μM) with 1 μl nuclease-free water prior to addition of 5 μl cDNA in BrightWhite real-time PCR fast 96-well plates (PrimerDesign, Chandler's Ford, United Kingdom). The plates were subsequently heated to 95°C for 20 s, followed by 40 cycles of 95°C for 1 s and 55°C for 20 s and melting-curve analysis at 95°C for 15 s, 60°C for 60 s, and 95°C for 15 s in a QuantStudio 6 Flex real-time PCR system with QuantStudio real-time PCR software (ThermoFisher Scientific, Loughborough, UK). Relative TNF-α and IL-1β gene expression was calculated with the β-actin gene as the reference gene and uninfected samples as the control using the method of Livak and Schmittgen (46).

View this table:
TABLE 2

Sequences of primers used for qPCR analysis

Primer specificity was ensured by the presence of single melting-curve peaks (see Fig. S9 in the supplemental material) and by running products on agarose gels, as previously described, to confirm single bands and correct product lengths (see Fig. S10 in the supplemental material). Primer efficiency was between 90 and 110% for all the primers used (see Fig. S11 in the supplemental material) and was determined using total rat RNA converted to cDNA, as previously described, and serially diluted 1:4 in nuclease-free water. Reference gene validation was performed by comparing gene stabilities across all the samples using NormFinder software (47). The β-actin gene was found to be the most stable reference gene (see Fig. S12 in the supplemental material).

TNF-α and IL-1β immunohistochemistry.Immunohistochemical staining of the tooth slices for TNF-α and IL-1β was performed based on methods used by Smith et al. (48). Rat lung was used as a positive control for TNF-α and IL-1β following fixation in 10% (wt/vol) neutral buffered formalin at room temperature for 24 h; dehydration through a series of 50% (vol/vol), 70% (vol/vol), 95% (vol/vol), and 100% (vol/vol) ethanol, followed by 100% (vol/vol) xylene, for 5 min each; and embedding in paraffin wax. The paraffin-embedded tooth slices and lung samples were cut, using a microtome, into 5-μm sections and incubated on glass slides at 65°C for 1 h. The samples were subsequently rehydrated through a series of xylene and 100% (vol/vol), 95% (vol/vol), and 70% (vol/vol) ethanol and double-distilled water for 5 min each. Endogenous peroxidase activity within the tissue sections was quenched by incubation in 3% (wt/vol) hydrogen peroxide for 10 min, followed by 2 washes for 2 min in Tris-buffered saline (TBS). Nonspecific binding was blocked with 3% (vol/vol) normal horse serum (Vector Laboratories, Peterborough, UK) in TBS for 30 min. The sections were incubated for 1 h with primary antibodies for TNF-α and IL-1β (Santa Cruz Biotechnology, Heidelberg, Germany) diluted 1:50 in TBS containing 1% (wt/vol) bovine serum albumin (Sigma-Aldrich, Gillingham, UK). An immunoreactivity assay was then performed using a Vectastain ABC peroxidase detection kit (Vector Laboratories, Peterborough, UK). Negative controls included omission of the primary antibody and replacement of the primary antibody with immunoglobulin G isotype diluted to the working concentration of the primary antibody. The sections were counterstained with 0.05% light green for 30 s, dehydrated with 100% ethanol and xylene for 10 min each, and mounted using VectaMount permanent mounting medium (Vector Laboratories, Peterborough, UK) prior to imaging using a Nikon digital camera and ACT-1 imaging software (Nikon UK Ltd., Surrey, UK).

SDS-PAGE of bacterial proteins.An overnight culture of S. anginosus and E. faecalis in BHI broth was prepared and diluted to 102 CFU/ml. S. anginosus and E. faecalis were cultured at 37°C and 5% CO2 for 24 h alone or in combination at ratios of 50:50 and 90:10. The suspensions were centrifuged at 5,000 × g for 5 min, and the supernatant was used for analysis of supernatant proteins. The pellet was lysed in radioimmunoprecipitation assay (RIPA) buffer by vortexing for 30 s followed by 30 s ultrasonication at 50 J using a Branson (Connecticut, USA) SLPe sonifier. Protein concentrations in the supernatant and the bacterial pellet were quantified using a bicinchoninic acid (BCA) assay (ThermoFisher Scientific, Loughborough, UK) and 20 μg of protein in Laemmli buffer (Bio-Rad, Hertfordshire, UK) separated by SDS-PAGE at 200 V for 40 min. Gels were stained using a Bio-Rad Silver Stain Plus kit according to the manufacturer's instructions and imaged using a Gel Doc EZ System (Bio-Rad, Hertfordshire, UK).

E. faecalis supernatant and heat-killed E. faecalis treatments.An overnight culture of E. faecalis was diluted in 20 ml DMEM-BHI medium to give a starting inoculum of 102 CFU/ml, as previously described. After incubation for an additional 24 h at 37°C and 5% CO2, the suspension was centrifuged at 5,000 × g for 5 min. The supernatant was filtered through a 0.22-μm syringe filter and frozen overnight at −20°C before freeze-drying for 24 h using a ScanVac CoolSafe freeze-dryer (LaboGene, Lynge, Denmark). The pellet of bacteria was resuspended in 20 ml of PBS and centrifuged at 5,000 × g for 5 min. This step was repeated to ensure minimal carryover of the culture supernatant. The pellet was then resuspended in 20 ml DMEM-BHI and heated to 100°C for 1 h. The solution was then frozen overnight at −20°C before freeze-drying as previously described. Twenty milliliters of sterile DMEM-BHI was also frozen and freeze-dried as a control. All freeze-dried samples were individually resuspended in 20 ml of sterile DMEM-BHI and used to culture rat tooth slices for RT-qPCR of cytokines and immunohistochemistry of TNF-α and IL-1β, as previously described.

Statistical analysis.A one-way analysis of variance (ANOVA) was performed using the data analysis package in Excel (Microsoft, Reading, UK) to determine the relative significance of the differences between the infected groups and the controls in terms of cell counts, bacterial coverage, and cytokine expression. The Tukey-Kramer test was used in conjunction with ANOVA to compare the significant differences between all possible pairs of means. A P value of ≤0.05 was considered significant.

ACKNOWLEDGMENT

This work was supported by the Dunhill Medical Trust (grant number R232/1111).

FOOTNOTES

    • Received 17 December 2017.
    • Returned for modification 19 January 2018.
    • Accepted 21 February 2018.
    • Accepted manuscript posted online 26 February 2018.

  • Supplemental material for this article may be found at https://doi.org/10.1128/IAI.00871-17.

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KEYWORDS

coculture
Enterococcus faecalis
ex vivo
host
infection
model
polymicrobial
pulpal infection
pulpitis
Streptococcus anginosus

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