In Vitro Anti-Biofilm and Antibacterial Properties of Streptococcus downii sp. nov.

microorganisms Article In Vitro Anti-Biofilm and Antibacterial Properties of Streptococcus downii sp. nov. Maigualida Cuenca 1,†, María Carmen Sánchez 1,*,† , Pedro Diz 2, Lucía Martínez-Lamas 3, Maximiliano Álvarez 3, Jacobo Limeres 2, Mariano Sanz 1and David Herrera 1   Citation: Cuenca, M.; Sánchez, M.C.; Diz, P.; Martínez-Lamas, L.; Álvarez, M.; Limeres, J.; Sanz, M.; Herrera, D. In Vitro Anti-Biofilm and Antibacterial Properties of Streptococcus downii sp. nov.. Microorganisms 2021,9, 450. https:// doi.org/10.3390/microorganisms 9020450 Academic Editor: Georgios N. Belibasakis Received: 21 January 2021 Accepted: 19 February 2021 Published: 22 February 2021 Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations. Copyright: © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/). 1ETEP (Etiology and Therapy of Periodontal and Peri-Implant Diseases) Research Group, University Complutense of Madrid (UCM), 28040 Madrid, Spain; [email protected] (M.C.); [email protected] (M.S.); [email protected] (D.H.) 2Medical-Surgical Dentistry Research Group (OMEQUI), Health Research Institute of Santiago de Compostela (IDIS), University of Santiago de Compostela (USC), 15705 Santiago de Compostela, Spain; [email protected] (P.D.); [email protected] (J.L.) 3Clinical Microbiology, Microbiology and Infectology Group, Galicia Sur Health Research Institute, Hospital Álvaro Cunqueiro, Complejo Hospitalario Universitario de Vigo, Vigo, 36312 Galicia, Spain; [email protected] (L.M.-L.); [email protected] (M.Á.) *Correspondence: [email protected]; Tel.: +34-913-941-674 † These authors equally contributed to this work. Abstract: The aim of this study was to evaluate the potential anti-biofilm and antibacterial activities of Streptococcus downii sp. nov. To test anti-biofilm properties, Streptococcus mutans, Actinomyces naeslundii, Veillonella parvula, Fusobacterium nucleatum, Porphyromonas gingivalis, and Aggregatibacter actinomycetemcomitans were grown in a biofilm model in the presence or not of S. downii sp. nov. for up to 120 h. For the potential antibacterial activity, 24 h-biofilms were exposed to S. downii sp. nov for 24 and 48 h. Biofilms structures and bacterial viability were studied by microscopy, and the effect in bacterial load by quantitative polymerase chain reaction. A generalized linear model was constructed, and results were considered as statistically significant at p< 0.05. The presence of S. downii sp. nov. during biofilm development did not affect the structure of the community, but an anti-biofilm effect against S. mutans was observed (p< 0.001, after 96 and 120 h). For antibacterial activity, after 24 h of exposure to S. downii sp. nov., counts of S. mutans (p= 0.019) and A. actinomycetemcomitans ( p= 0.020 ) were significantly reduced in well-structured biofilms. Although moderate, anti-biofilm and antibacterial activities of S. downii sp. nov. against oral bacteria, including some periodontal pathogens, were demonstrated in an in vitro biofilm model. Keywords: oral biofilm; Streptococcus downii sp. nov.; caries; Streptococcus mutans; periodontal diseases; Aggregatibacter actinomycetemcomitans 1. Introduction The oral cavity has one of the richest microbial communities within the human microbiome, with around 200 cultivable bacterial species and approximately 1000 phylotypes detected by 16S rRNA gene sequencing [ 1 , 2 ]. In healthy subjects, the majority of these species are commensal and maintain a symbiotic relation with the host, being the predominant phyla: Firmicutes, Proteobacteria, Actinobacteria, Bacteroidetes, and Fusobacteria [ 3 ]. Among Firmicutes the most abundant species belongs to Streptococcus, Veillonella, and Lactobacillus genera [ 4 ]. However, under specific environmental conditions, existing pathobionts may overgrow and become pathogenic, mainly by disturbing the homeostasis between the bacterial challenge and the host immune responses (dysbiosis) [ 5 ], which can also be associated with changes in microbial metabolism and/or shifts in bacterial diversity. Moreover, within the oral cavity, overgrow of microorganisms usually occurs forming highly structured poly-microbial communities, as biofilms, what may hinder the efficacy of host defenses and natural antimicrobial strategies. Microorganisms 2021,9, 450. https://doi.org/10.3390/microorganisms9020450 https://www.mdpi.com/journal/microorganisms Microorganisms 2021,9, 450 2 of 16 Intervention studies have confirmed the etiological importance of biofilms by demonstrating that mechanical biofilm disruption is a key step in the effective management of biofilm-related oral conditions, caries, and periodontal diseases. In the treatment of the most aggressive/severe cases of periodontitis, more effective outcomes have been demonstrated when mechanical debridement has been supplemented with the adjunctive use of antibiotics [ 6 ] and antiseptics [ 7 ]. The use of antimicrobials, however, is controversial, due to the possible occurrence of secondary effects and the development of bacterial resistances [6–9], which calls for exploring alternative strategies. One of these alternative strategies is to foster a health-associated microbiota that will transform the dysbiotic state into re-establishing homeostasis. One strategy in this direction has been the use of live microorganisms (probiotics), mainly bifidobacteria and lactobacilli, which have shown adjunctive benefits in the prevention and treatment of periodontal diseases [ 10 – 20 ]. This benefit has been attributed to the ability of these probiotics (i) to occupy a habitat, and thus reduce pathogen adhesion and colonization; (ii) to boost the host immune response; and (iii) since they have inherent antimicrobial activity [ 21 ]. Another strategy, seldom explored and investigated, has been the use of indigenous oral species, which will occupy and ecological niche, and thus will reduce the adhesion and growth of opportunistic pathogens and pathogenic species [ 22 – 26 ]. Streptococcus spp., such as Streptococcus sanguinis,Streptococcus cristatus,Streptococcus salivarius, or Streptococcus mitis, have shown their ability to inhibit the in vitro colonization of epithelial cells by Aggregatibacter actinomycetemcomitans [ 24 , 25 ], S. mitis showed antagonism in the adhesion of Porphyromonas gingivalis [ 23 ], and Streptococcus dentisani, isolated from caries-free individuals, showed growth inhibition of periodontal pathogens and against pathogens implicated in dental root infections, in pure culture [ 22 , 27 , 28 ]. Moreover, S. dentisani, attached to gingival cells in vitro , inhibits periodontal pathogens by competition, adherence, and displacement mechanisms [ 28 ]. Similarly, bifidobacteria may have the capacity of suppressing the growth of P. gingivalis by reducing key nutritional factor(s) in the environment [26]. Recently, derived from the analysis of the oral microbiota of patients with Down syndrome a novel species of bacteria of the Streptococcus oralis group, Streptococcus downii sp. nov., has been described [ 29 ]. By inhibition assays, S. downii sp. nov. has exhibited a potentially antimicrobial effect against the cariogenic bacteria Streptococcus mutans and against the periodontal pathogens Veillonella parvula and A. actinomycetemcomitans [29]. However, despite the fact that oral bacteria are organized in biofilms, most of these in vitro studies cited have investigated this antibacterial potential against bacteria in the planktonic state, rather than assessing this effect on biofilm models, thus mimicking a situation closer to what happens in vivo . Therefore, based on the results of individual species—species that determined the specific effect on S. mutans,V. parvula and A. actinomycetemcomitans, the purpose of this investigation was to assess (i) the potential anti-biofilm activity of S. downii sp. nov., either by inhibiting the growth of selected oral bacteria and/or by interfering with biofilm formation when growing on an in vitro biofilm model, and (ii) the potential antibacterial effects against oral bacteria in mature biofilms. 2. Materials and Methods 2.1. Isolates, Culturing, and Bacterial Growth Conditions S. downii sp. nov. strain CECT 9732T, isolated from a supragingival dental biofilm sample of an individual with Down syndrome [ 29 ], and the reference bacterial strains S. mutans ATCC 25175, V. parvula NCTC 11810, Actinomyces naeslundii ATCC 19039, F. nucleatum DMSZ 20482, A. actinomycetemcomitans DSMZ 8324, and P. gingivalis ATCC 33277 were used to develop a multi-species biofilm model. Bacteria were cultured anaerobically (10% H 2 , 10% CO 2 , and balance N 2 ) on blood agar plates (Blood Agar Oxoid No 2; Oxoid, Basingstoke, UK), supplemented with 5% (v/v) sterile horse blood (Oxoid, Basingstoke, UK), 5.0 mg mL −1 hemin (Sigma-Aldrich, St. Louis, MO, USA) and 1.0 mg mL -1 menadione (Merck, Darmstadt, Germany) for 72 h at 37 ◦C. Microorganisms 2021,9, 450 3 of 16 2.2. Anti-Biofilm Activity of S. downii sp. nov. in an in Vitro Biofilm Model Figure 1shows the experimental design followed for the study of the anti-biofilm properties of S. downii sp. nov. strain CECT 9732T against bacteria in an oral biofilm model. Microorganisms 2021, 9, x FOR PEER REVIEW 3 of 16 Basingstoke, UK), supplemented with 5% (v/v) sterile horse blood (Oxoid, Basingstoke, UK), 5.0 mg mL −1 hemin (Sigma-Aldrich, St. Louis, MO, USA) and 1.0 mg mL -1 menadione (Merck, Darmstadt, Germany) for 72 h at 37 °C. 2.2. Anti-Biofilm Activity of S. downii sp. nov. in an in Vitro Biofilm Model Figure 1 shows the experimental design followed for the study of the anti-biofilm properties of S. downii sp. nov. strain CECT 9732T against bacteria in an oral biofilm model. Figure 1. Scheme of the antibacterial assays carried out in the study. For abbreviations, see the text. 2.2.1. Biofilm Development Biofilms were developed on hydroxyapatite (HA) discs in a static biofilm model, based on the model described by Sánchez et al. [30] with some modifications. In brief, representative colonies of all species were selected randomly and were grown on brainheart infusion (BHI) medium (Becton, Dickinson and Company, Franklin Lakes, NJ, USA) supplemented with 2.5 g L −1 mucin (Oxoid, Basingstoke, UK), 1.0 g L −1 yeast extract (Oxoid, Basingstoke, UK), 0.1 g L −1 cysteine (Sigma-Aldrich, St. Louis, MO, USA), 2.0 g L −1 sodium bicarbonate (Merck, Darmstadt, Germany ), 5.0 mg L −1 hemin (Sigma-Aldrich, St. Louis, MO, USA), and 1.0 mg L −1 menadione (Merck, Darmstadt, Germany) and 0.25% (v/v) glutamic acid (Sigma-Aldrich, St. Louis, MO, USA), in anaerobic condition at 37 °C for 24 h. The bacterial growth was harvested at late exponential phase (measured by spectrophotometry), and a mixed bacterial suspensions in supplemented BHI medium were prepared in order to develop biofilms, using different starting concentrations, depending on their growth rate: 10 3 colony forming units (CFU) mL −1 of S. mutans, 10 5 CFU mL −1 of V. parvula and A. naeslundii, and 10 6 CFU mL −1 of F. nucleatum, A. actinomycetemcomitans, and P. gingivalis. Sterile HA discs [7-mm diameter and 1.8 (standard deviation—SD = 0.2) mm thickness (Clarkson Chromatography Products, Williamsport, PA, USA)] were placed in a multi-well tissue culture plate (Greiner Bio-one, Frickenhausen, Germany). Each well was inoculated with 1.5 mL of the mixed bacterial suspension, and to assess the anti-biofilm Figure 1. Scheme of the antibacterial assays carried out in the study. For abbreviations, see the text. 2.2.1. Biofilm Development Biofilms were developed on hydroxyapatite (HA) discs in a static biofilm model, based on the model described by Sánchez et al. [ 30 ] with some modifications. In brief, representative colonies of all species were selected randomly and were grown on brainheart infusion (BHI) medium (Becton, Dickinson and Company, Franklin Lakes, NJ, USA) supplemented with 2.5 g L −1 mucin (Oxoid, Basingstoke, UK), 1.0 g L −1 yeast extract (Oxoid, Basingstoke, UK), 0.1 g L −1 cysteine (Sigma-Aldrich, St. Louis, MO, USA), 2.0 g L −1 sodium bicarbonate (Merck, Darmstadt, Germany ), 5.0 mg L −1 hemin (Sigma-Aldrich, St. Louis, MO, USA), and 1.0 mg L −1 menadione (Merck, Darmstadt, Germany) and 0.25% (v/v) glutamic acid (Sigma-Aldrich, St. Louis, MO, USA), in anaerobic condition at 37 ◦ C for 24 h. The bacterial growth was harvested at late exponential phase (measured by spectrophotometry), and a mixed bacterial suspensions in supplemented BHI medium were prepared in order to develop biofilms, using different starting concentrations, depending on their growth rate: 10 3 colony forming units (CFU) mL −1 of S. mutans, 10 5 CFU mL −1 of V. parvula and A. naeslundii, and 10 6 CFU mL −1 of F. nucleatum,A. actinomycetemcomitans, and P. gingivalis. Sterile HA discs [7-mm diameter and 1.8 (standard deviation—SD = 0.2) mm thickness (Clarkson Chromatography Products, Williamsport, PA, USA)] were placed in a multiwell tissue culture plate (Greiner Bio-one, Frickenhausen, Germany). Each well was inoculated with 1.5 mL of the mixed bacterial suspension, and to assess the anti-biofilm activity, treated wells were inoculated with 10 3 CFU mL −1 of S. downii sp. nov. final concentration (recovered at late-exponential phase by centrifugation of an overnight culture and resuspended in fresh modified BHI medium). Control biofilms, not exposed to S. downii sp. nov., and treated ones were then incubated in anaerobiosis at 37 ◦ C, for 12, 24, 48, 72, 96, and 120 h. Plates only containing culture media were also incubated to check for sterility. Three independent trials (on three different occasions) were carried out. Microorganisms 2021,9, 450 4 of 16 2.2.2. DNA Isolation and Quantitative Polymerase Chain Reaction (qPCR) Before the DNA isolation, discs were sequentially rinsed in 2 mL of sterile buffer saline (PBS) (immersion time per rinse, 10 s), three times, in order to remove non-adherent bacteria. Biofilms were then disrupted by vortex for 2 min in 1 mL of PBS. The DNA from the biofilm was extracted using an ATP Genomic DNA Mini Kit ® (ATP biotech. Taipei, Taiwan) according to the manufacturer’s recommendations. The bacterial 16S rRNA gene sequence was amplified by qPCR, according to the hydrolysis probes 5 ´ nuclease method (Table 1). The qPCR amplification was performed in a total reaction mixture volume of 10 µ L. The reaction mixtures contained 5 µ L of 2 × master mixture (LC 480 Probes Master; Roche, Mannheim, Germany), optimal concentrations of primers and probes (300, 300 and 300 nM, respectively, for S. mutans,Streptococcus spp., A. naeslundii and P. gingivalis; 750, 750, and 400 nM for V. parvula; 300, 300, and 200 nM for A. actinomycetemcomitans; and, finally, 600, 600, and 300 nM for F. nucleatum) and 2 µ L of DNA from samples. The negative control was 2 µ L of sterile water (Water PCR grade, Roche Mannheim, Germany). The samples were subjected to an initial amplification cycle of 95 ◦ C for 10 min, followed by 40 cycles at 95 ◦ C for 15 s and 60 ◦ C for 1 min. Analyses was performed with a LightCycler ® 480 II thermocycler (Roche Mannheim, Germany). LightCycler ® 480 Multiwell Plates 384 and sealing foils were used (Roche Mannheim, Germany). Each DNA sample was analyzed in duplicate. Table 1. Primers and probes used for quantification of genomic DNA from the target bacteria. Primers and probes were targeted against 16S rRNA gene (Obtained from Life Technologies Invitrogen (Carlsbad, CA, USA) and Roche (Roche Diagnostic GmbH; Mannheim, Germany)). Bacteria Sequence (50–30) Length (bp) Streptococcus spp. Forward CAACGATACATAGCCGACCTGAG 97 Reverse TCCATTGCCGAAGATTCC Probe 6FAM-CTCCTACGGGAGGCAGCAGTAGGGA-BBQ S. mutans Forward GCCTACAGCTCAGAGATGCTATTCT 58 Reverse GCCATACACCACTCATGAATTGA Probe 6FAM-TGGAAATGACGGTCGCCGTTATGAA-TMR V. parvula Forward TGCTAATACCGCATACGATCTAACC 66 Reverse GCTTATAAATAGAGGCCACCTTTCA Probe 6FAM-CTATCCTCGA+T+GCC+GA-BBQ A. naeslundii Forward GGCTGCGATACCGTGAGG 103 Reverse TCTGCGATTACTAGCGACTCC Probe 6FAM-CCCTAAAAGCCGGTCTCAGTTCGGAT-BBQ P. gingivalis. Forward GCGCTCAACGTTCAGCC 67 Reverse CACGAATTCCGCCTGC Probe 6FAM-CACTGAACTCAAGCCCGGCAGTTTCAA-TAMRA A. actinomycetemcomitans Forward GAACCTTACCTACTCTTGACATCCGAA 80 Reverse TGCAGCACCTGTCTCAAAGC Probe 6FAM-AGAACTCAGAGATGGGTTTGTGCCTTAGGG-TAMRA F. nucleatum Forward GGATTTATTGGGCGTAAAGC 162 Reverse GGCATTCCTACAAATATCTACGAA Probe 6FAM-CTCTACACTTGTAGTTCCG-TAMRA Quantification of bacteria by qPCR was based on standard curve. The correlation between Cq values and CFU mL −1 was automatically generated through the software (LC 480 Software 1.5; Roche Mannheim, Germany). Since the primers and probes targeting Microorganisms 2021,9, 450 5 of 16 Streptococcus spp. detected all Streptococcus present in the sample, the number of S. downii sp. nov. was calculated by subtracting the number of S. mutans. 2.2.3. Analysis of Biofilms by Confocal Laser Scanning Microscopy (CLSM) Biofilms grown on HA discs were stained with LIVE/DEAD ® BacLight TM Bacterial Viability Kit solution (Molecular Probes B. V., Leiden, The Netherlands) at room temperature, with 1:1 fluorocromes ratio, in the dark for 10 min (SD = 1). The obtained fully hydrated biofilms were studied with a confocal laser scanning microscope, using a fixed-stage Ix83 Olympus inverted microscope, coupled to an Olympus FV1200 confocal system (Olympus; Shinjuku, Tokyo, Japan) with a × 63 water-immersion lenses (Olympus, Shinjuku, Tokyo, Japan). At least three separate and representative locations were selected from the HA discs covered with biofilm. With the use of a dedicated software (Olympus ® software (Olympus, Shinjuku, Tokyo, Japan) and Image analysis FIJI ® software (Image J v. 2.0.0-rc-65 /1.52b) a z-series of scans (xyz) of 0.5 µm thickness (8 bits, 1024 ×1024 pixels) were analyzed. 2.2.4. Analysis of Biofilms by Scanning Electron Microscope (SEM) Specimens were fixed in a solution at 4% paraformaldehyde and 2.5% glutaraldehyde for 4 h, at 4 ◦ C. The discs were washed once in phosphate buffer saline (PBS) and another time in sterile water (immersion time per washed 10 min) and then dehydrated through a series of graded ethanol solutions (30, 50, 70, 80, 90, and 100%; immersion time per series 10 min). After that, specimens were critical point dried, sputter-coated with gold and analysed by electron microscopy JSM 6400 (JSM6400; JEOL, Tokyo, Japan) with a back-scattered electron detector and an image resolution of 25 kV. 2.3. Antibacterial Effect of S. downii sp. nov. Against Oral Species in an Already Established in Vitro Biofilm Figure 1presents the experimental design of the study on the antibacterial activity of S. downii sp. nov. strain CECT 9732T against bacteria, in an in vitro oral biofilm model. 2.3.1. Biofilm Development Biofilms were developed as described previously (Section 2.2.1), and a mixed bacterial suspensions in supplemented BHI medium were prepared in order to develop biofilms, containing 10 3 CFU mL −1 of S. mutans, 10 5 CFU mL −1 of V. parvula and A. naeslundii, and 10 6 CFU mL -1 of F. nucleatum,A. actinomycetemcomitans, and P. gingivalis. HA discs (Clarkson Chromatography Products, Williamsport, PA, USA) and 1.5 mL of the bacterial suspension were placed in the multi-well tissue culture plate (Greiner Bio-one, Frickenhausen, Germany) and incubated at 37 ◦ C in anaerobiosis for 24 h. After that, the 24 h developed biofilms were: (i) exposed to 1 mL of S. downii sp. nov. at 10 8 CFU mL −1 in supplemented BHI medium or (ii) in the case of controls, exposed to 1 mL of fresh supplemented BHI medium. Biofilms were then incubated for additional 24 and 48 h at 37 ◦C in anaerobiosis. Three independent trials (on three different occasions) were carried out. 2.3.2. Biofilm Analysis The 24 h biofilms and the biofilms re-incubated another 24–48 h in the presence or not (controls) of S. downii sp. nov., were analysed. Confocal laser scanning microscopy and scanning electron microscopy (Sections 2.2.3 and 2.2.4) displayed the biofilms structures and bacterial viability. Quantitative polymerase chain reaction was used to assess the effect of S. downii sp. nov. in bacterial load (amounts of each bacterium expressed as CFU mL −1 ; Section 2.2.2). 2.4. Statistical Analysis The primary outcome variable was the count of viable bacteria present in the in vitro developed biofilms, expressed as viable CFU mL −1 of S. downii sp. nov., S. mutans, V. parvula,A. naeslundii,A. actinomycetemcomitans,P. gingivalis, and F. nucleatum. An Microorganisms 2021,9, 450 6 of 16 experiment-level analysis was performed for each parameter of the study (n= 3). Shapiro– Wilk goodness-of-fit tests and distribution of data were used to assess normality. Data were expressed as means and standard deviations (SD). In order to evaluate the impact of S. downii sp. nov. and the time of biofilmdevelopment (up to 120 h) and their interaction with the primary outcome variable (counts expressed in CFU mL −1 ), a general linear model was constructed for each bacterial species, using the method of maximum likelihood and Bonferroni corrections for multiple comparisons. A p-value < 0.05 was considered statistically significant. A software package (IBM SPSS Statistics 21.0; IBM Corporation, Armonk, NY, USA) was used for all data analysis. 3. Results 3.1. Anti-Biofilm Activity of S. downii sp. nov. in an in Vitro Biofilm Model The impact of S. downii sp. nov. on the obtained multispecies biofilms was assessed from 12 to 120 h. Figure 2depicts the kinetic profiles of the six bacterial species used in this in vitro biofilm model (strains S. mutans,A. naeslundii,V. parvula,F. nucleatum, P. gingivalis , and A. actinomycetemcomitans), quantified by qPCR, when forming biofilms with or without the presence of S. downii sp. nov. In biofilms exposed to S. downii sp. nov., it could be detected this bacterial species incorporated in the biofilm as early as 12 h of biofilm evolution and up to 120 h, with an average concentration of 2.7 × 10 7 CFU mL −1 (SD = 1.5 ×107). In both obtained biofilms, with or without the presence of S. downii sp. nov, the six bacterial species were incorporated already after 12 h of incubation, and were present for up to 120 h. Only the growth of S. mutans was significantly different when S. downii sp. nov. was present in the biofilms (Figure 2). This impact was not observed in the early stages of biofilm formation, since at 12 h, both biofilms were similar (p> 0.05). From 24 to 72 h there was a clear trend, but without being statistically significant (p> 0.05 in all cases). However, differences were evident and statistically significant in the stationary phase (96 and 120 h) of the biofilm formation (p< 0.001, in both cases) (Figure 2). The kinetics of the rest of initial colonizers (V. parvula and A. naeslundii), and that of the periodontal pathogens ( F. nucleatum ,A. actinomycetemcomitans, and P. gingivalis) was not affected by the presence of S. downii sp. nov. during the 120 h of incubation (p> 0.05 in all cases). Biofilm structure and bacterial viability were studied by CLSM. During the initial growth phase, and after 24 h of incubation, biofilms containing S. downii sp. nov. depicted a well-structured bacterial community, similar to biofilms without S. downii sp. nov. ( Figure 3a,b ), with a live/dead ratio of 2.75 (SD = 0.2) for biofilms without S. downii sp. nov. and 2.4 (SD = 0.8) for biofilms containing S. downii sp. nov. This tendency was maintained during the exponential phase of biofilm development and, after 72 h, both modalities of biofilms showed a similar architecture, corresponding to a mature bacterial community (Figure 3c,d), with a live/dead ratio of 3.0 (SD = 0.1), for biofilms without S. downii sp. nov., and 2.7 (SD = 0.2), for biofilms containing S. downii sp. nov. However, and in agreement with the previously described qPCR data, in the stationary phase of biofilms (from 96 to 120 h, Figure 3e,f), it could be observed a higher percentage of dead cells in biofilms containing S. downii sp. nov., when compared with biofilms without S. downii sp. nov. The live/dead ratio was 2.0 (SD = 0.1) for biofilms without S. downii sp. nov. and 1.4 (SD = 0.4) for biofilms containing S. downii sp. nov. Microorganisms 2021,9, 450 7 of 16 Microorganisms 2021, 9, x FOR PEER REVIEW 7 of 16 Figure 2. Kinetics of incorporation of the six selected bacterial strains in the biofilm [expressed as logarithm of Colony Forming Units per mL, (log CFU mL −1 )] on the two biofilm modalities compared in the study: control biofilms (composed of Streptococcus mutans, Veillonella parvula, Actinomyces naeslundii, Fusobacterium nucleatum, Aggregatibacter actinomycetemcomitans, and Porphyromonas gingivalis) and experimental biofilms, incorporating also Streptococcus downii sp. nov. Analyses have been performed with quantitative polymerase chain reaction, in biofilms from 12 h to 120 h of incubation, using specific primers and probes directed to the 16S rRNA gene. *p < 0.005. Biofilm structure and bacterial viability were studied by CLSM. During the initial growth phase, and after 24 h of incubation, biofilms containing S. downii sp. nov. depicted a well-structured bacterial community, similar to biofilms without S. downii sp. nov. (Figure 3a,b), with a live/dead ratio of 2.75 (SD = 0.2) for biofilms without S. downii sp. nov. and 2.4 (SD = 0.8) for biofilms containing S. downii sp. nov. This tendency was maintained during the exponential phase of biofilm development and, after 72 h, both modalities of biofilms showed a similar architecture, corresponding to a mature bacterial community (Figure 3c,d), with a live/dead ratio of 3.0 (SD = 0.1), for biofilms without S. downii sp. nov., and 2.7 (SD = 0.2), for biofilms containing S. downii sp. nov. However, and in agreement with the previously described qPCR data, in the stationary phase of biofilms (from 96 to 120 h, Figure 3e,f) , it could be observed a higher percentage of dead cells in biofilms containing S. downii sp. nov., when compared with biofilms without S. downii sp. nov. The live/dead ratio was 2.0 (SD = 0.1) for biofilms without S. downii sp. nov. and 1.4 (SD = 0.4) for biofilms containing S. downii sp. nov. Figure 2. Kinetics of incorporation of the six selected bacterial strains in the biofilm [expressed as logarithm of Colony Forming Units per mL, (log CFU mL −1 )] on the two biofilm modalities compared in the study: control biofilms (composed of Streptococcus mutans, Veillonella parvula, Actinomyces naeslundii, Fusobacterium nucleatum, Aggregatibacter actinomycetemcomitans, and Porphyromonas gingivalis) and experimental biofilms, incorporating also Streptococcus downii sp. nov. Analyses have been performed with quantitative polymerase chain reaction, in biofilms from 12 h to 120 h of incubation, using specific primers and probes directed to the 16S rRNA gene. * p< 0.005. Microorganisms 2021,9, 450 8 of 16 Microorganisms 2021, 9, x FOR PEER REVIEW 8 of 16 Figure 3. Confocal micrographs that represented a 2D maximum projection of the series along fixed axis of the control biofilms (a,c,e), composed by Streptococcus mutans, Veillonella parvula, Actinomyces naeslundii, Fusobacterium nucleatum, Aggregatibacter actinomycetemcomitans, and Porphyromonas gingivalis, and experimental biofilms, incorporating also Streptococcus downii sp. nov. (b,d,f), after 24 h (a,b), 72 h (c,d) and 120 h (e,f) of growth. LIVE/DEAD® BacLightTM Bacterial Viability Kit stain was used to assess the vitality of cells (live cells in green and dead cells in red color; yellowish corresponded to damage cells but still alive). The analysis by SEM showed similar findings (Figure 4). After 120 h or growth, it would seem that the proportion of S. mutans forming chains of cocci was reduced in biofilms containing S. downii sp. nov., compared to grown biofilms without S. downii sp. nov. (Figure 4e,f). Figure 3. Confocal micrographs that represented a 2D maximum projection of the series along fixed axis of the control biofilms ( a , c , e ), composed by Streptococcus mutans, Veillonella parvula, Actinomyces naeslundii, Fusobacterium nucleatum, Aggregatibacter actinomycetemcomitans, and Porphyromonas gingivalis, and experimental biofilms, incorporating also Streptococcus downii sp. nov. ( b , d , f ), after 24 h ( a , b ), 72 h ( c , d ) and 120 h ( e , f ) of growth. LIVE/DEAD ® BacLight TM Bacterial Viability Kit stain was used to assess the vitality of cells (live cells in green and dead cells in red color; yellowish corresponded to damage cells but still alive). The analysis by SEM showed similar findings (Figure 4). After 120 h or growth, it would seem that the proportion of S. mutans forming chains of cocci was reduced in biofilms containing S. downii sp. nov., compared to grown biofilms without S. downii sp. nov. (Figure 4e,f). Microorganisms 2021,9, 450 9 of 16 Microorganisms 2021, 9, x FOR PEER REVIEW 9 of 16 Figure 4. Scanning electron microscope images of the control biofilms (a,c,e), composed by Streptococcus mutans, Veillonella parvula, Actinomyces naeslundii, Fusobacterium nucleatum, Aggregatibacter actinomycetemcomitans and Porphyromonas gingivalis, and experimental biofilms, incorporating also Streptococcus downii sp. nov. (b,d,f), after 24 h (a,b), 72 h (c,d) and 120 h (e,f) of growth. A similar architecture of biofilms can be observed, in both presence and absence of S. downii sp. nov., with biofilms covering the disc surfaces with flat homogenous layers of cells, combined with bacterial clusters, showing channels inside the structure. 3.2. Antibacterial Effect of S. downii sp. nov. Against Oral Species in an Already Established in Vitro Biofilm After 24 h of incubation, well-structured biofilms were developed. The presence of the six inoculated bacterial species was confirmed by qPCR, and their live/dead ratio, measured by CLSM, was 2.0 (SD = 0.1) (Figure 5). Figure 4. Scanning electron microscope images of the control biofilms ( a , c , e ), composed by Streptococcus mutans, Veillonella parvula, Actinomyces naeslundii, Fusobacterium nucleatum, Aggregatibacter actinomycetemcomitans and Porphyromonas gingivalis, and experimental biofilms, incorporating also Streptococcus downii sp. nov. ( b , d , f ), after 24 h ( a , b ), 72 h ( c , d ) and 120 h ( e , f ) of growth. A similar architecture of biofilms can be observed, in both presence and absence of S. downii sp. nov., with biofilms covering the disc surfaces with flat homogenous layers of cells, combined with bacterial clusters, showing channels inside the structure. 3.2. Antibacterial Effect of S. downii sp. nov. Against Oral Species in an Already Established in Vitro Biofilm After 24 h of incubation, well-structured biofilms were developed. The presence of the six inoculated bacterial species was confirmed by qPCR, and their live/dead ratio, measured by CLSM, was 2.0 (SD = 0.1) (Figure 5). These biofilms were then exposed to 10 8 CFU mL −1 of S. downii sp. nov. for 24 and 48 h and examined by CLSM (Figure 6). The obtained 24 h biofilms showed similar well-structured bacterial communities when compared those non-exposed versus exposed to S. downii sp. nov. (Figure 6a,b, respectively) and with live/dead ratio of 3.0 (SD = 0.3) and 4.2 (SD = 1.7), respectively. In those exposed to S. downii sp. nov., a thicker live biomass could be observed, possibly caused by the additional bacteria incorporated (Figure 6a,b). After 48 h of exposition, in spite of having a similar biofilm structure, the percentage of dead bacteria was significantly higher in the biofilms exposed to S. downii sp. nov. Microorganisms 2021,9, 450 16 of 16 44. Lee, D.K.; Park, S.Y.; An, H.M.; Kim, J.R.; Kim, M.J.; Lee, S.W.; Cha, M.K.; Kim, S.A.; Chung, M.J.; Lee, K.O.; et al. Antimicrobial activity of Bifidobacterium spp. isolated from healthy adult Koreans against cariogenic microflora. Arch. Oral. Biol. 2011 ,56, 1047–1054. [CrossRef] 45. Schwendicke, F.; Horb, K.; Kneist, S.; Dorfer, C.; Paris, S. Effects of heat-inactivated Bifidobacterium BB12 on cariogenicity of Streptococcus mutans in vitro. Arch. Oral Biol. 2014,59, 1384–1390. [CrossRef] 46. Suzuki, N.; Yoneda, M.; Hatano, Y.; Iwamoto, T.; Masuo, Y.; Hirofuji, T. Enterococcus faecium WB2000 Inhibits Biofilm Formation by Oral Cariogenic Streptococci. Int. J. Dent. 2011,2011, 834151. [CrossRef] 47. Soderling, E.M.; Marttinen, A.M.; Haukioja, A.L. Probiotic lactobacilli interfere with Streptococcus mutans biofilm formation in vitro. Curr. Microbiol. 2011,62, 618–622. [CrossRef] [PubMed] 48. Marttinen, A.M.; Haukioja, A.L.; Keskin, M.; Soderling, E.M. Effects of Lactobacillus reuteri PTA 5289 and L. paracasei DSMZ16671 on the adhesion and biofilm formation of Streptococcus mutans. Curr. Microbiol. 2013,67, 193–199. [CrossRef] 49. Lin, X.; Chen, X.; Chen, Y.; Jiang, W.; Chen, H. The effect of five probiotic lactobacilli strains on the growth and biofilm formation of Streptococcus mutans. Oral Dis. 2015,21, e128–e134. [CrossRef] 50. Teughels, W.; Loozen, G.; Quirynen, M. Do probiotics offer opportunities to manipulate the periodontal oral microbiota? J. Clin. Periodontol. 2011,38, 159–177. [CrossRef] [PubMed] 51. Van Essche, M.; Quirynen, M.; Sliepen, I.; Van Eldere, J.; Teughels, W. Bdellovibrio bacteriovorus attacks Aggregatibacter actinomycetemcomitans. J. Dent. Res. 2009,88, 182–186. [CrossRef] [PubMed] 52. Loozen, G.; Boon, N.; Pauwels, M.; Slomka, V.; Rodrigues Herrero, E.; Quirynen, M.; Teughels, W. Effect of Bdellovibrio bacteriovorus HD100 on multispecies oral communities. Anaerobe 2015,35, 45–53. [CrossRef] [PubMed] 53. Cuenca, M.; Marín, M.J.; Nóvoa, L.; O‘Connor, A.; Sánchez, M.C.; Blanco, J.; Limeres, J.; Sanz, M.; Diz, P.; Herrera, D. Periodontal Condition and Subgingival Microbiota Characterization in Subjects with Down Syndrome. Appl. Sci. 2021,11, 778. [CrossRef] 54. Novoa, L.; Sanchez, M.D.C.; Blanco, J.; Limeres, J.; Cuenca, M.; Marin, M.J.; Sanz, M.; Herrera, D.; Diz, P. The Subgingival Microbiome in Patients with Down Syndrome and Periodontitis. J. Clin. Med. 2020,9, 2482. [CrossRef] 55. Fernandez, M.; Hudson, J.A.; Korpela, R.; de los Reyes-Gavilan, C.G. Impact on human health of microorganisms present in fermented dairy products: An overview. Biomed. Res. Int. 2015,2015, 412714. [CrossRef]