The study was performed in accordance with the Helsinki Declaration of 2013. Informed consent to the study procedure was obtained from all participants. The protocol was approved by the Bioethics Committee at the Jagiellonian University in Krakow (No. 122.6120.99.2016). Informed consent was obtained from all individual parents.

The study was conducted between 2015 and 2016 and included 143 pediatric patients from the Department of Pediatric Dentistry, Institute of Dentistry, Jagiellonian University in Krakow, Poland. The study included bacterial strains isolated from dental plaque of patients (n = 143; average age 4.6 ± 0.76) diagnosed with early childhood caries (ECC) of the deciduous teeth. The results marked with reference numbers were entered onto a standardized examination chart. Examinations were conducted based on the criteria established by the World Health Organization for epidemiological studies (WHO, 1997).

Children were divided into two subgroups: subjects with initial stage decay, defined as the non-cavitated group (1–2 in ICDAS codes); and subjects with extensive decay, defined as the cavitated group (5–6 in ICDAS II codes) (International Caries Detection and Assessment System Coordinating Committee, 2012). Ninety-seven patients were qualified to the non-cavitated group (47 girls, aged 4.68 ± 0.91 years and 50 boys, aged 4.6 ± 0.61). Forty six patients were assigned to the cavitated group (20 girls, aged 4.75 ± 0.85 years and 26 boys, aged 4.35 ± 0.63).

The exclusion criteria were as follows: age below 2 or above 6 years, diabetes, periodontal disease, epithelial dysplasia, and inflammatory lesions of the oral mucosa. Antibiotics, non-steroid, anti-inflammatory medications, corticosteroids, and vitamin intake within the last 3 months also resulted in a patient’s exclusion from the study. Dental plaque was determined using the OHI-S index (Simplified Oral Hygiene Index) (Wei and Lang, 1982).

After classification of the patients for the research, supragingival plaque (pooled from all surfaces of individual primary first teeth) samples were collected using sterile dental explorers. Each subject rinsed their mouth for 1 min with distilled water, before plaque collection. Material was collected from fasted participants in the morning (between 8 AM and 10 AM), before toothbrushing and any clinical examination. The collected plaque was put into sterile tubes with 0.5 ml phosphate buffered saline (PBS), inhibiting O₂, and carried on ice to the lab within 1 h. Plaque samples were then sonicated for 30 s, vortexed to disperse and to obtain a homogeneous suspension and clarified by centrifugation at 2000 × g for 10 min at 4°C. A total of 50 μl aliquots of clarified supernatants were used for traditional plate culturing methods on HLR-S medium [HL Ritz medium containing 40 g tryptic soy agar (TSA), 20% sucrose, 0.3 U/ml bacitracin, 1.75 μg/ml polymyxin B sulfate and 0.5 μg/ml crystal violet] for three 10-fold dilutions of each plaque sample. All plates were incubated under 85% N₂, 10% CO₂, 5% O₂ conditions at 37°C for 48 h. Samples with colony counts of more than 130 (10⁴ cells/ml) were considered positive.

After obtaining pure colony samples on the HLR-S medium, single colonies were inoculated on the surface of TSA plates with 5% sheep blood and incubated under previously mentioned conditions. Then, the characteristics of individual colonies were evaluated, including the colonial shape and form, the type of hemolysis, and the Gram-staining.

Phenotypic identification of the isolated S. mutans species was performed using a commercial biochemistry identification test: STREPTOtest24 (Lachema, Pliva).

Biotyping from selected enzymatic reactions of the above commercial test was performed for S. mutans as the dominant bacteria species. Two approaches to biotyping criteria based on enzymatic activity profiles were used. The first approach consisted of the selection of appropriate enzymes (INU, MLB, TGT) based on the analysis of their activity in the population of the tested strains.

The second method for biotype determination consisted of the use of an unattended (without a priori available knowledge) statistical data analysis. Data were divided into groups (clusters), so that each group was as homogenous as possible (strains within the group were similar) and at the same time, clusters differed from one another (strains from different groups had the lowest possible number of common characters).

Single pure S. mutans colonies were grown in 4 ml of BHI broth (Difco, BD, USA) supplemented with 5% sucrose for 8 h. Cells were harvested in the logarithmic phase of growth and washed three times with 40 mM potassium phosphate buffer (pH 7.0).

In the growth of S. mutans were analyzed by using a flow cytometer (LSRII, BD Biosciences Immunocytometry Systems, San Jose, CA, USA). Aggregates and doublets were excluded using a gating strategy related to the width versus height of the forward scatter (FSC) and side scatter (SSC).

A standardized suspension of the tested microorganisms was prepared in PBS to give 1 on the McFarland scale using a MicroSpek dual DSM densitometer. The density of the inoculum [1 × 10⁷ colony forming units (CFUs)/ml] was confirmed by counting single colonies after 24 h growth under the same conditions as described for BHI agar for S. mutans.

The capacity for biofilm formation was evaluated using a closed model based on a microtiter plate. The biomass of molded/non-degraded biofilm was measured by staining with crystal violet (CV) according to the method described by Peeters et al. (2008). At the outset, 100 μl of a standardized S. mutans suspension was transferred to each well of a 96-well microtiter plate, which was incubated for 90 min to start the initial adhesion of S. mutans. The suspension was aspirated and each well was washed three times with PBS to remove planktonic cells. Subsequently, 100 μl of fetal bovine serum was added to each well and incubated under the same conditions for 8 h. The wells were washed three times in PBS and 100 μl of a standardized S. mutans suspension was added to each well. Control samples were 100 μl of PBS (PBS control) or 100 ml of BHI broth with 5% sucrose (medium control).

To initiate biofilm growth and its further development, 60 μl of BHI with 5% sucrose was added to each well. The medium was changed once and the plates were incubated for 48 h.

After 4, 6, 8, 10, 12, 14, 16, 18, 20, 22, and 24 h of incubation, the cells were washed twice with PBS. After addition of PBS, the biofilm was broken by homogenization in an ultrasonic homogenizer (Hielscher UP50H) for 30 s at an amplitude of 25%. Subsequently, serial dilutions of the solution were prepared by inoculating MSBS medium (containing bacitracin and sucrose) with 100 μl of the bacterial suspension and incubated for 48 h. The number of colonies (CFU/ml) was determined. The test was repeated in the two experiments at different time points, with a number of repeats in groups.

Biofilm biomass was evaluated after 4, 6, 8, 10, 12, 14, 16, 18, and 24 h of incubation by staining with (CV). The biofilm was maintained in 99% methanol for 20 min, and air dried. Subsequently, 125 μl of 0.1% CV solution was added for 20 min, and the wells were rinsed three times with PBS. After each washing step, plates were dried by shaking. After the last washing step, the samples were allowed to dry thoroughly. Finally, bounded CV was released by adding 200 μl of 95% ethanol, followed by incubation for 15 min at room temperature. The contents of the wells were mixed by repeated pipetting and then 125 μl of the suspension was transferred to the wells of a clean 96-well flat-bottom microtiter plate. The absorbance was measured at a wavelength of λ_max = 540 nm. All steps were performed at room temperature. The study was conducted in two independent experiments. A biofilm formation curve was plotted based on the obtained data.

Cover glasses (Agar Scientific, UK) of a 13 mm diameter were placed in the wells of a 24-well plate for biofilm formation in accordance with the procedure described above. At 4, 6, 8, 10, 12, 14, 16, 18, 20, 22, and 24 h, the samples were fixed in 1 ml of 2.5% glutaraldehyde solution for 1 h, dehydrated in a graded ethanol series for 20 min each, followed by immersion in 100% alcohol for 1 h. The disks were dried in an incubator for 24 h. After drying, the sample was transferred into aluminum sections and sprayed with gold (160 s, 40 mA). The samples were examined and photographed under a JEOL JSM-35CF scanning electron microscope (SEM) at 20–25 kV in the SEM Laboratory of the Otolaryngology Clinic, University Hospital in Krakow, Poland. These experiments were performed at each time point on three cultures of mono-species biofilm.

Pyruvate kinase activity was determined using S. mutans cells after 4, 6, 8, 10, 12, 14, 16, 18, 20, 22, and 24 h of a single-species biofilm formation using a continuous assay coupled with lactate dehydrogenase (LDH), in which the change in absorbance at 340 nm due to oxidation of NADH was measured using a POLARstar Omega microplate spectrophotometer (BMG Labtech, Germany).

Obtained cells were inoculated into 396 ml of sterile 40 mM potassium phosphate buffer (pH = 7.0) containing 10 mM beta-mercaptoethanol (BME), and grown for 24 h. Subsequently, cells were disrupted by sonic oscillation (15 min, 0°C, 200 W, 2A, Hielscher UP50H oscillator). The cell extract was centrifuged (17,500 × g for 30 min at 4°C) and the supernatant was used for further purification of the enzyme. All subsequent procedures were performed on ice.

Cleaning efficiency is ∼140-fold and recovery amounted to ∼10%. The purified enzyme was stored at -20°C in TRIS buffer with 10% glycerol until further enzymatic activity analysis.

The reaction for PK activity measurement contained 1 mM ADP, 1 mM phosphoenolpyruvate (PEP), 0.4 mM glucose-6-phosphate (G6P), 0.12 mM NADH (Santa Cruz Biotechnology, USA),10 μg/ml LDH (Carl Roth, Germany), 100 mM KCl, 10 mM MgCl₂, and 0.13 μg/ml of the tested sample containing PK in 67 mM Tris-HCl buffer (pH 7.0) in a total volume of 250 μl. The activity was determined spectrophotometrically by recording the change in absorbance at 340 nm (μmol/min/mg of protein). PK activity was defined as the quantity of PK which catalyzed the formation of one micromole of PEP per minute. The reaction was carried out for 10 min at 30°C on a POLARstar Omega microplate spectrophotometer (BMG Labtech, Germany). Values of kinetic parameters such as K_m and V_max were calculated with respect to the change in the substrate (PEP) concentration (0–2.0 mM) at a fixed ADP concentration of 1 mM fitting in GraphPad Prism 7 (GraphPad TM Software, USA).

Statistical analysis was performed using R 3.2.3 (R Development Core Team, 2009). The normality of the data was assessed by the Shapiro–Wilk test and, where appropriate, non-parametric analyses were performed. Data are expressed as median ± upper and lower quartiles. Strains were divided into clusters using agglomerative hierarchical clustering approach with common setting: complete linkage and Ward metric. To determine the relationship between PK, CFU, and the optical density (OD) and the stage of disease and biotypes, the Kruskal–Wallis test was used. To answer the question as to how exactly this dependence looks in particular biotypes, a post hoc analysis was made (Dunn’s test). Markers were ascertained using the Spearman coefficient of correlation, analysis of covariance (ANCOVA) was performed to assess the connections between PK, CFU and OD, biotypes and the forms of the disease, corrected for the effect of confounding factors (age, sex). A value of p < 0.05 was considered statistically significant.

In this project a total of 143 strains of S. mutans were isolated from supragingival plaque. Two approaches were used in determining the biotyping criteria based on enzymatic activity profiles of STREPTOtest 24.

The first approach was to select appropriate enzymes based on an analysis of their activity in the tested strain population. Three enzymes were selected, whose incidence among the tested strains was in a quartile deviation between 15 and 85%. These are inulin – 40.56% melibiose – 83.22%, and tagatose – 21.68%. The other 21 enzymes occurring in the test (STREPTOtest24) were rejected, because they did not meet the established criteria for enzyme inclusion (i.e., a frequency or enzyme activity between 15 and 85% in the test population).

The acceptance of these enzymes for biotyping criteria allowed the definition of eight enzymatic profiles, conventionally denoted by consecutive letters of the alphabet from A to H, as consistently shown: profile A occurring in 0.70% of the strains was characterized by a lack of activity of inulin (INU), melibiose (MLB), and tagatose (TGT). Profile B occurring in 6.99% of the strains showed tagatose activity and inactive inulin and melibiose. Profile C occurring in 42.66% of the strains was characterized by melibiose activity and inactive inulin and tagatose. Profile D with inactive inulin and active melibiose and tagatose occurred in 9.09% of the strains. Profile E with inactive melibiose and tagatose, and active inulin occurred in 7.69% of the strains. Profile F was characterized by a lack of melibiose activity, active inulin and tagatose, and occurred in 1.40% of the strains. Profile G was characterized by a lack of tagatose activity, active inulin and melibiose and occurred in 27.27% of the strains. Profile H was characterized by the activity of all three enzymes (inulin, melibiose, and tagatose) and occurred in less than 4.20% of the strains. The most numerously represented profile is ‘C,’ characterized by a lack of inulin or tagatose activity.

The distribution of the obtained profiles was heterogeneous; therefore, strains whose profiles differed only according to MLB were grouped into one biotype (MLB was the nearest to being excluded from the range 15–85%). The following four biotypes are given in four Roman numerals: I–IV. Biotype I, with a frequency of 43.36% (n = 62), contained profiles A and C, and showed melibiose activity without active inulin and tagatose. Biotype II (16.08%, n = 23) consisted of profiles B and D and showed tagatose activity, and inactive inulin with partially active melibiose. Biotype III (34.97%, n = 50) consisted of profiles E and G, and included strains with active inulin, inactive tagatose and partial melibiose activity. Biotype IV (5.59%, n = 8) consisted of profiles F and H and was characterized by partial melibiose activity and active inulin and tagatose.

Another way to determine biotypes was the application of statistical unattended (without a priori available knowledge) data analysis.

The results of the division into clusters are shown in dendrograms. Strains were differentiated based on the activity of melibiose and tagatose, which enabled the extraction of three biotypes – the main branches of the diagram (Figure 1) marked with the colors red, green and blue as A, B, C. The figure also indicates the distribution of enzymatic profiles obtained using the arbitrary method; red cluster is biotype II; green is biotype I, and blue is biotype III.

While performing analysis in terms of the severity of the decay of four biotypes obtained in the arbitrary method, it was noted that the analyzed groups differed significantly (p = 0.003; Fisher’s exact test) (Figures 1A,B). The strains belonging to biotype I in the non-cavitated group constituted 49.48% (n = 48) and 30.43% (n = 14) in the cavitated group. The strains classified to biotype II constituted 11.34% (n = 11) in the non-cavitated group and 26.09% (n = 12) in the cavitated one. In turn, strains from biotype III were 37.11% (n = 36) in the non-cavitated and 30.43% (n = 14) in the cavitated group. In the case of biotype IV, strains from the non-cavitated group were less than 2.06% (n = 2) and 13.04% (n = 6) in the cavitated group.

While performing clustering in terms of the severity of decay of three clusters from the most well-fitted dendrogram, it was noted that the analyzed groups differed significantly (p < 0.001; Fisher’s exact test) (Figures 1A,C). Strains classified in cluster A in the non-cavitated group represented 0% (n = 0) and 26.09% (n = 12) in the cavitated group. In cluster B, 56.70% (n = 55) of the strains were non-cavitated and 34.78% (n = 16) belonged to the cavitated group. In cluster C, 43.30% (n = 42) of the strains belonged to the non-cavitated group while 39.13% (n = 18) to the cavitated one.

Logs of the number of microorganisms (S. mutans) (CFU/ml) forming a biofilm at the respective time points are statistically significant between both groups. p-value is less than 0.05 for all time points (4–24 h); therefore, the analyzed groups differed significantly in terms of the amount of microorganisms at any time point. At each of the time points of single-species biofilm formation, the amount of S. mutans in the –cavitated– group was significantly higher than in the –non-cavitated– one.

The highest number of microorganisms forming a biofilm was observed after 18 h in strains isolated from dental plaque from the cavitated group of children compared with non-cavitated children (7.67 ± 7.65–7.69 vs. 7.54 ± 7.52–7.57, p < 0.001; U Mann–Whitney test).

The number of microorganisms (S. mutans) (log CFU/ml) forming a biofilm at the respective time points was not dependent on sex (p > 0.05 for all time points, U Mann–Whitney test).

The total biomasses of microorganisms (S. mutans) forming a biofilm at the respective time points are statistically significant. p-value is less than 0.05 for all time points (4–24 h); therefore, the analyzed groups differed significantly in terms of the amount of microorganisms at any time point. At each of the time points of single-species biofilm formation, the amount of S. mutans in the cavitated group was significantly higher than in the non-cavitated one.

The highest absorbance (the greatest amount of microorganisms cumulated in the biofilm) was observed after 18 h in strains isolated from dental plaque from the cavitated group of children compared with non-cavitated children (0.13 vs. 0.12, p < 0.001; U Mann–Whitney test).

The total biomass of microorganisms (S. mutans) forming biofilm at the particular time points did not depend on sex (p > 0.05 for all time points; U Mann–Whitney test).

Attention was paid to the relationship between the number of biofilm-forming microorganisms (S. mutans) (log CFU/ml) and their biomass (OD) at the respective time points. The correlation is statistically significant (p < 0.05) at each time point of biofilm formation. These relationships are positive: the higher the log (CFU/ml), the higher the OD, and vice versa. The strongest relationship was noted after 18 h of biofilm formation (coefficient of correlation r = 0.71, p < 0.001) (Figure 2).

In vitro biofilm formation was also analyzed using a SEM, where mature single-species biofilms were observed after 18 h of incubation (Figures 3, 4). Figures 3, 4 shows the S. mutans cells, extracellular matrix (ECM) and water channels responsible for nutrition within the biofilm. The stages of biofilm formation are clearly identifiable: from the initial phase of the microorganism adhesion to the surface of a flat polystyrene; biofilm maturation when an exopolysaccharide matrix is created and other bacteria species join the biofilm; and finally, the formation of a mature structure and a slowdown in the growth tempo of individual bacteria.

The correlation between the log (CFU/ml) and PK activity is statistically significant at each time point of single-species biofilm formation (p < 0.05). The strongest correlation between the measured parameters was observed after 18 h of biofilm formation, r = 0.771, p < 0.001; Spearman correlation coefficient (Figure 5). These relationships are positive: the higher the log (CFU/ml), the higher the PK activity, and vice versa.

Similarly, attention has been paid to the relationship between the PK activity at particular time points and the biomass of a formed biofilm (OD = 540 nm). These relationships are positive: the higher the PK activity, the higher the OD, and vice versa. This time, correlation between the measured parameters in a mature biofilm (after 18 h) was strong, r = 0.771, p < 0.001; Spearman correlation coefficient (Figure 6).

At each time point of biofilm formation, the PK activity in the cavitated group was significantly higher than in the non-cavitated group (p < 0.05 for all time points; Mann–Whitney Test). Differences at particular time points were as follows: 1.16 ± 1.14–1.21 (median and Q₁–Q₃ quartiles) after 4 h for the non-cavitated group (n = 97) and 1.24 ± 1.23–1.25 for the cavitated one (n = 46). After 6 h: 1.18 ± 1.16–1.23 for the non-cavitated and 1.26 ± 1.24–1.27 for the cavitated group. After 8 h the median values were: 1.2 ± 1.18–1.25 in the non-cavitated and 1.28 ± 1.26–1.29 in the cavitated group. At subsequent points of time, PK activity increased steadily, peaking at 18 h of biofilm formation and equaling 1.36 ± 1.33–1.4 in the non-cavitated group and 1.56 ± 1.52–1.6 in the cavitated one. After 20 h of biofilm formation, PK activity consistently decreased, giving after 24 h median values of 1.26 ± 1.24–1.3 in the non-cavitated and 1.46 ± 1.38–1.48 in the cavitated group (p < 0.05, Mann–Whitney test).

Four analyzed biotypes statistically differed in terms of the activities of PK at each of the time points of biofilm formation (p < 0.05, Kruskal–Wallis test). After 4, 6, 8, 10, 12, and 14 h, PK activities in biotypes I and II were significantly higher than in biotype III (p = 0.014, p = 0.019, p = 0.012, p = 0.004, p = 0.004, and p = 0.011, respectively; post hoc Dunn’s test, Kruskal–Wallis test). After 16 h, the PK activity in biotype II was significantly higher than in biotype III (p = 0.014; post hoc Dunn’s test, Kruskal–Wallis test). After 18 h, the activities of PK in biotypes I, II, and IV were significantly higher than in biotype III (p = 0.002; post hoc Dunn’s test, Kruskal–Wallis test). After 20 and 22 h, the activities of PK in biotypes II and IV were significantly higher than in biotype III (p = 0.004 for both time points; post hoc Dunn’s test, Kruskal–Wallis test). After 24 h, the PK activity in biotype IV was significantly higher than in biotype III (p = 0.013; post hoc Dunn’s test, Kruskal–Wallis test) (Table 1).

Three analyzed clusters statistically differed in terms of the activities of PK at each of the time points of biofilm formation (p < 0.05, Kruskal–Wallis test). At each of the time points, PK activity in cluster A was significantly higher than in the other (B, C) clusters (p ≤ 0.001 for all time points; post hoc Dunn’s test, Kruskal–Wallis test) (Table 2).

The analyzed biotypes significantly differed in terms of the amounts of microorganisms forming a biofilm CFU/ml for eight time points (p < 0.05, Kruskal–Wallis test). Post hoc analysis showed that the number of microorganisms contained in the biofilm after 18 h in cluster A was significantly higher than in clusters B and C (in terms of OD and CFU differentiation, division into three clusters is better) (p = 0.006; post hoc Dunn’s test). Taking into account the differences in log CFU/ml between four biotypes (from arbitrary method), a great diversity was observed. Post hoc analysis (Kruskal–Wallis test + Dunn’s test; p < 0.05) showed that after 4 and 6 h CFU/ml in biotype IV was significantly higher than in biotype I (p = 0.284 and p = 0.081; post hoc Dunn’s test); after 14 h, CFU/ml in biotype IV was significantly higher than in biotype III (p = 0.049; post hoc Dunn’s test); after 16 h, CFU/ml in biotype IV was significantly higher than in biotypes I and III (p = 0.026; post hoc Dunn’s test); after 18 and 20 h, CFU/ml in biotype II was significantly higher than in biotype III (p = 0.017; post hoc Dunn’s test); and after 20 and 24 h, CFU/ml in biotype II was significantly higher than in biotypes III and I (p = 0.037 and p = 0.022; post hoc Dunn’s test) (Tables 3, 4).

The analyzed biotypes were significantly different in terms of the biomass of a formed biofilm (OD) for eight time points (p < 0.05, Kruskal–Wallis test). Post hoc analysis revealed that the biofilm biomass after 18 h in cluster A was significantly higher than in clusters B and C (p < 0.05, Dunn’s test).

Taking into account the differences in OD between the four biotypes (from the arbitrary method) a great diversity can be observed. Post hoc analysis (Kruskal–Wallis test + Dunn’s test; p < 0.05) showed that after 6 h, OD in biotype II was significantly higher than in biotype I; after 8, 10, 14, 18, and 20 h, OD in biotype II was significantly higher than in biotypes I and III; after 16 and 22 h, OD in biotype II was significantly higher than in biotype III.

Considering the division into three clusters (clusterization method), these results were more consistent than those from the arbitrary method. Adoption of the above-mentioned criterion in completing the division into three clusters seems to be more reasonable (in the context of OD and CFU/ml differentiation), as in that case the same relationship can be observed at each time point of biofilm formation; while for four biotypes (arbitrary method) this relationship is quite strongly “mixed” (Tables 5, 6).