Table of Contents Author Guidelines Submit a Manuscript
The Scientific World Journal
Volume 2014 (2014), Article ID 165201, 6 pages
http://dx.doi.org/10.1155/2014/165201
Research Article

Genotypic Diversity and Virulence Traits of Streptococcus mutans Isolated from Carious Dentin after Partial Caries Removal and Sealing

Department of Social and Preventive Dentistry, Faculty of Dentistry, Federal University of Rio Grande do Sul, Ramiro Barcelos 2492, 90035-003 Porto Alegre, RS, Brazil

Received 29 August 2013; Accepted 13 November 2013; Published 21 January 2014

Academic Editors: P. G. Arduino, L. N. Baratieri, and S. Barros

Copyright © 2014 Nailê Damé-Teixeira et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract

The aim of this study was to compare the genotypic diversity and virulence traits of Streptococcus mutans isolated from carious dentin before and after partial dentin caries removal (PDR) and sealing. Carious dentin samples were obtained three months before and after the PDR and cavity sealing. Up to seven isolates of each morphological type of S. mutans were selected and strain identity was confirmed using gtfB primer. Genotyping was performed by arbitrary primer-PCR (AP-PCR). Acidogenesis and acidurance of the genotypes were evaluated as virulence traits. A paired -test and a Wilcoxon test were used to compare the virulence of genotypes. A total of 48 representative S. mutans isolates were genotyped (31 before and 17 after the sealing). At least one of the genotypes found before the sealing was also found on dentin after the sealing. The number of genotypes found before the sealing ranged from 2 to 3 and after the sealing from 1 to 2 genotypes. No difference was observed in the acidogenesis and acidurance between genotypes isolated before and after the sealing. In conclusion, genotypic diversity of S. mutans decreased after the PDR and sealing, but the virulence traits of S. mutans remained unchangeable.

1. Introduction

Stepwise excavation has been an alternative treatment for deep caries lesions since the conventional treatment based on complete dentin caries removal could generate pulp exposure and poor dental prognosis [1]. The stepwise excavation involves the partial removal of the decayed tissue, temporary sealing, reopening of the cavity, and the complete removal of the carious tissue followed by restoration [13]. After the sealing period, the filling is removed and complete caries removal is performed [2, 4, 5]. Partial carious dentin removal (PDR) in one session, keeping a layer of carious dentin beneath restoration, has been proposed as an alternative approach to the stepwise excavation, avoiding failures due to the loss of temporary filling, additional costs, discomfort to the patient, and the possibility of pulp exposure during the second reopening procedure [6, 7]. This treatment has shown success in clinical studies over time without the need of cavity reentry [68]. However, the persistence of viable bacteria in dentin after PDR has raised doubts regarding the long-term effectiveness of this treatment [9].

After the sealing period, a limited supply of nutrients is left for the bacteria that survive underneath the restoration. Strains that are fit for physical, chemical, biological, and environmental changes might dominate and get established after the sealing. Paddick et al. [10] showed that only those bacteria capable of producing the enzymes required for the cleavage of the terminal sugars from the glycoprotein were recovered from the dentin after the cavity sealing. The low nutrient supply underneath the restorations could lead to a modification of the residual biofilm.

Although a limited number of microorganisms persist under restorations a few months after the PDR and sealing [2, 3, 11, 12], some cariogenic bacteria may be found within the remaining microorganisms, such as mutans streptococci, which is currently found on sealed carious dentin [1017]. These bacteria are capable of producing acids due to fermentation of dietary carbohydrates (acidogenesis) and surviving in that low-pH environment (acidurance), which makes them an important cariogenic microorganism related to caries initiation and progression [18]. However, the relationship between the residual S. mutans found on carious dentin beneath restoration after the PDR and caries progression is still unclear. Thus, it is important to better understand how the environmental changes induced by the dentin sealing affect the diversity and virulence traits of the remaining mutans streptococci.

In this context, genotyping methods based on arbitrarily primed PCR (AP-PCR) have revealed that the community of S. mutans isolated from saliva and dental plaque is diverse [19, 20]. Additionally, it has been shown that distinct S. mutans genotypes may exhibit distinct cariogenic potential [21, 22]. Therefore, the aim of this study was to compare the genotypic diversity of S. mutans isolated from carious dentin before and after the PDR and sealing. We hypothesize that S. mutans genotypes are selectively found after the PDR and sealing and exhibit low cariogenic potential.

2. Methods

2.1. Origin of the Samples

The samples were derived from a previous clinical trial. Briefly, patients () with permanent molars with carious lesions located in the middle third of the dentin were selected [23]. The patients were submitted to the PDR and sealing with a biocompatible material for 3 months. Dentin samples were obtained after the PDR (before and after the sealing), by a sterile bur, transferred to and diluted in reduced transport fluid, and plated on Mitis Salivarius Bacitracin (MSB) agar (Difco Laboratories, Detroit, MI, USA). Up to 7 isolates of each morphological type found in these cultures were selected and analyzed based on colony morphology. After subculture, each isolate was stored in Brain-Heart Infusion (BHI) (HiMedia, Mumbai, India) with 15% (v/v) glycerol at −20°C for further analysis. From 18 patients, only 4 patients presented S. mutans isolates before and after the sealing.

2.2. Extraction of Genomic DNA

Streptococcus mutans were grown from frozen stocks on BHI agar and incubated for 24 h at 37°C in microaerophilic conditions. The genomic DNA was extracted from colonies resuspending them in 50 μL of sterile ultrapure water [24]. PCR with species-specific primers to gtfB (-ACTACACTTTCGGGTGGCTTGG- and -CAGTATAAGCGCCAGTTTCATC-) was performed to confirm the identity of S. mutans isolates [19] (Invitrogen, SG, Milanese, Italy). The PCR amplifications were performed with 50 μL total volume, including 1 μL of the target DNA, 0.25 μL of Taq DNA polymerase (5 U/μL), 5 μL of 10 × PCR buffer, 2.5 μL of 50 mM MgCl2, 1 μL of deoxynucleoside triphosphate mix (10 mM), and 1 μL of each primer (10 mM). The amplifications were performed under the following conditions: 30 cycles of 95°C for 30 s, 59°C for 30 s, and 72°C for 1 min, followed by 1 cycle of 95°C for 30 s, 59°C for 30 s, and a final extension of 72°C for 5 min. Genomic DNA of S. mutans strain UA159 (provided by FIOCRUZ, Rio de Janeiro, Brazil) and ultrapure water were applied in all the PCR baths, as positive and negative controls. The PCR products were analyzed by electrophoresis on 2% agarose gel and stained with SYBR Green 1.6%, at 100 V for 45 min. Bands were visualized under UV illumination. All the chemicals were provided by Invitrogen (SG, Milanese, Italy).

2.3. Genotypic Analysis of S. mutans Isolates by AP-PCR

AP-PCR assays were performed with the arbitrary primer OPA 02 (-TGCCGAGCTG-) [19]. The DNA amplification occurred under the following conditions: 95°C for 2 min, for initial denaturation, and 45 cycles of 94°C for 30 s, 36°C for 30 s, and 72°C for 1 min, with a final extension at 72°C for 5 min. Streptococcus mutans strain UA159 and ultrapure water were used as controls. Products of AP-PCR were analyzed by electrophoresis on 1% agarose gel and stained with SYBR Green 1.6%, at 96 V for 4 hours.

2.4. Virulence Factors Analysis

The virulence traits of genotypes were evaluated as described elsewhere [19]. Frozen stocks of each S. mutans genotype were regrown on BHI agar plates and incubated at 37°C for 24 h. Two loops of 1 μL were inoculated into 30 mL of BHI broth supplemented with 1% glucose and incubated at 37°C for 18 h. In order to evaluate the ability of S. mutans genotypes to lower the suspension pH through glycolysis, 10 mL of the microbial suspension was centrifuged and resuspended in 50 mM KCl/1 mM MgCl2 (Synth, São Paulo, Brazil). The pH of the solution was adjusted to around 7.0, and glucose was added to a final concentration of 55.5 mM. The pH was monitored for 180 min using a glass electrode previously calibrated with pH standards (pH 4.0 and 6.8). The area above the curve (AAC) was determined considering pH 6.5 as a cutoff point. The experiment was performed in duplicate and S. mutans UA 159 was used as a control in all the tests.

The ability of S. mutans genotypes to tolerate acidic environments was evaluated using 10 mL of the overnight growth suspension (described above) for 18 h in BHI broth/glucose. This suspension was diluted to 1 : 20 in BHI/glucose, and the growth was monitored until OD550 = 0.5. The suspensions were then centrifuged, and the pellets were washed once with 0.1 M glycine buffer (pH 7.0) (Nuclear, São Paulo, Brazil). The washed pellets were then resuspended in 0.1 M glycine buffer pH 2.8, 5.0, and 7.0. Immediately after the resuspension (time zero) and after 30 min and 60 min of incubation at 37°C, aliquots were serially diluted in phosphate buffer (pH 7.2), plated on BHI agar, and incubated at 37°C for 48 h. Cell viability at each time point was expressed as the percent of growth in relation to time zero.

2.5. Data Analysis

Images of AP-PCR fingerprints were captured by a digital camera (Canon Inc., Tokyo, Japan) and stored in Image File Format for visual analysis. For analysis of the S. mutans genotypic profiles from the same patient, AP-PCR products from the isolates obtained before and after the sealing were always resolved side by side in the same gel for visual comparisons. Thus, genotypic diversity was compared among isolates before and after the sealing samples. Isolates were considered as having the same genotypic identity when they presented identical AP-PCR product-size profiles. Later, a side-by-side analysis on the same gel was performed using each genotype (duplicate) from all the patients to verify the similarities between S. mutans isolates from different patients. Two blinded and calibrated examiners performed the visual analysis. Cohen’s Kappa value was 0.76. Double cases were discussed and a consensus was reached. Genotypes (number and proportion) were described in each patient before and after the sealing (descriptive analysis). Data were transformed to log10 due to data dispersion, and the normal distribution was confirmed using a histogram and Kolmogorov-Smirnov test, except for the pH 2.8 analysis. A paired -test and a Wilcoxon test were used to compare the virulence of genotypes (acidogenesis and acidurance) found before and after the sealing. The significance level was set at 5%. The virulence-traits data were analyzed by (1) the paired comparison between genotypes found before the sealing with the same genotypes found after the sealing (genotypes A, D, E, G, and I) and by (2) the comparison between the means of acidogenesis and acidurance of all the genotypes found before the sealing with all the genotypes found after the sealing. Statistical tests were performed in SPSS 18.0 for Windows (IBM SPSS Statistics).

2.6. Ethical Considerations

The protocol of the clinical trial was approved by the ethics committee of the Faculty of Dentistry from the Federal University of Rio Grande do Sul (process no. 19218). Informed and written consent was obtained from all the individuals. All the participants received the treatment for basic dental needs.

3. Results

A total of 48 S. mutans isolates were obtained from carious dentin corresponding to 31 found before and 17 found after the sealing (Table 1). All of them were identified as being S. mutans by species-specific PCR (Figure 1). The molecular weight fragments generated by AP-PCR from the studied patients range between 490 and 5000 bp generating 9–12 DNA bands. The genotypic diversity of S. mutans found in one patient (patient 1) is shown in Figure 2. The number of genotypes found per patient varies between 2-3 and 1-2 before and after the sealing, respectively. At least one of the genotypes found before the sealing was also found on dentin after the sealing in all the patients. Only one patient presented one new genotype after the sealing (patient 3). Two patients (1 and 4) showed reduction in the number of genotypes before and after the sealing. In another patient (patient 2), the same genotypes were found before and after the sealing, although their proportion was different, with predominance of genotype D (Table 1). For patient 3, only one of the genotypes identified before the sealing was found after the sealing. That genotype was the most prevalent, and it colonized the dentin in addition to a new genotype exclusively found after the sealing (Table 1). No similarities were found between S. mutans isolates from different patients.

tab1
Table 1: Number of isolates and genotypes (%) of S. mutans isolated per patient from carious dentin before and after sealing.
165201.fig.001
Figure 1: S. mutans identification by species-specific gtfB primer. Lanes 1 to 6: isolates of S. mutans; lanes 7 and 8 correspond to the positive control (S. mutans UA 159) and negative control (water), respectively; lane 9: 100-bp DNA ladder.
165201.fig.002
Figure 2: Electrophoresis gel image from AP-PCR-generated fingerprints using primer OPA-02 of S. mutans reference strain (UA159) and isolates from carious dentin before and after sealing. Lanes 1 to 15 correspond to the S. mutans isolates before sealing; lanes 16 to 20 correspond to the S. mutans isolates after sealing; lane 21: negative control (water); lane 22: 250-bp DNA ladder.

The results of the analysis of the virulence traits are described in Table 2 (acidogenesis) and Table 3 (acidurance). In the acidogenesis analysis, no difference was observed in the AAC neither when the comparisons were made considering genotypes found before and after the sealing within the same patient () nor when the comparisons were made considering all genotypes found before and after the sealing (). Regarding the acidurance, no difference was observed in the growth of the genotypes at different pH when the analysis was performed considering paired genotypes found before and after the sealing.

tab2
Table 2: Acidogenesis (AAC; pH 6.5 as cutoff point) of S. mutans genotypes isolated from carious dentin before and after sealing.
tab3
Table 3: Acidurance of S. mutans genotypes isolated from carious dentin before and after sealing.

4. Discussion

This study aimed to compare the genotypic diversity and the virulence traits of S. mutans in carious dentin after PDR before and after the sealing, using AP-PCR fingerprinting analysis. We observed a reduction in the number of S. mutans isolates found after the sealing (Table 1). This is consistent with a reduction in counts of viable cells on carious dentin found after the cavity sealing [23]. AP-PCR has been used to evaluate the genotypic profile of S. mutans from saliva, biofilm, tongue, and dentin [1921, 2530]. The validity of the AP-PCR technique in the genotypic identification of microorganisms is assured by several comparisons made with other genotyping techniques [20, 31, 32].

Previous studies have shown a decrease in bacterial diversity [10, 33] after the PDR and sealing. A shift in the bacterial genotypes has also been observed [10], but no studies evaluating the genotypic diversity of S. mutans were found. In the present study, an altered genotypic diversity was observed after sealing the carious dentin.

It has been discussed that the residual bacteria found after the sealing have limited access to external nutrients. They are fed mainly by glycoproteins provided by the pulp and that nutrient restriction exerts a selective pressure over the residual bacteria [10]. In that scenario, a metabolism shift has been found in bacteria isolated from carious dentin after the sealing [10]. It is also likely that components of dead bacteria, which did not survive after the sealing, may contribute as the source of nutrients to those bacteria that survived below the restorations [10]. Considering that our dentin samples were collected 3 months after the sealing, these findings might explain the altered genotype prevalence found within different patients after the dentin sealing (Table 1). Some genotypes (B and F) were the most prevalent in dentin before the sealing; however, they were not detected after the dentin sealing (Table 1). The undetected levels of these genotypes after the sealing might have enhanced the competitiveness of other genotypes (A and G) that were most prevalent after the sealing. In patient 3 (Table 1), there was a shift in the prevalence of genotypes found before and after the sealing. We believe that the reduction in the proportion of genotypes D and J has created a better condition for the growth of genotypes E and I. Interestingly, only genotype H was found on dentin samples after the sealing. It means that the genotype was below the detection limit of the microbiological method used [34] on dentin samples before the sealing. Moreover, some rare or transient genotypes in a single sample might be missed in a complex microbiota, which could explain the appearance of a new genotype only after the sealing [35]. The sealing of the cavity reduced the prevalence of genotype F, which might have created better conditions for the growth of genotypes G and H. In an open environment, such as a dental caries cavity, the microbiota of the biofilm above the dentin is continuously exposed to the dietary carbohydrate that provides nutrients for the growth of the diverse oral flora. However, the sealing of carious cavity was responsible for the selection of genotypes that were capable of surviving in the presence of low nutrient availability. Besides reduced availability of nutrients during the sealing, the relative simplicity and homogeneity of these nutrients significantly affect the microbiota surviving under the restorations [10, 33].

Additionally, microbiological studies have reported a significant decrease in the bacterial infection in the residual carious dentin after the cavity sealing [2, 3, 11, 12]. However, despite this microbial residual contamination found after the sealing, the carious dentin becomes harder and drier, both characteristics of inactive lesions [2]. Therefore, the reduction in the number of S. mutans genotypes found after the cavity sealing represents to some extent the low number of genotypes frequently found in caries-free individuals [25, 27, 28].

It remains unclear, though, whether the bacteria underneath restorations represent some danger to the longevity of restorations and if they are more virulent than the bacteria prior to the sealing. In this study, no statistical difference was found regarding the virulence traits of S. mutans isolated before and after the sealing. Even under a low-nutrient-availability condition, the role of specific phenotypic traits on the prevalence of these genotypes remains the same. That means those genotypes found after the sealing might be metabolically active if a source of external carbohydrates is provided, and, in addition to that, those genotypes are still capable of developing an acid-tolerance response to the acidic condition. Thus, it is important to point out the need for a perfectly sealed restoration.

Moreover, acidurance and acidogenesis of genotypes found before and after the sealing (A, D, E, G, and I) were not different which means that equal genotype presented equal phenotype. In contrast, several studies have shown that distinct S. mutans genotypes might show distinct acidurance [21, 22]. The results of the acidurance analysis in the present study suggest that the higher pH observed in sealed carious dentin [36] is due to the decrease in the number of these acidogenic bacteria and the limited access to nutrients [10] and not by their lower capacity of acid production.

Therefore, it is important to know if the biofilm that survived underneath the restorations remains potentially cariogenic [10, 37]. According to Takahashi and Nyvad [38], it is not only important to describe which bacteria are involved in caries, but it is also important to know what is their function [38]. Although S. mutans is one important cariogenic microorganism related to caries, only 4 out of 18 patients showed this microorganism after the sealing. Besides this, the S. mutans that survived presents similar virulence in comparison to the initial S. mutans.

In conclusion, genotypic diversity of S. mutans was reduced after PDR and sealing using AP-PCR fingerprinting analysis. Additionally, there was not any difference in acidurance and acidogenesis between genotypes found before and after the sealing. Genotypes found after the PDR and sealing have the same cariogenic potential of those found before the sealing.

Conflict of Interests

The authors declare that they have no proprietary, financial, professional, or other personal interests of any nature or kind in any product, service, and/or company that could be construed as influencing the position presented in, or the review of, the paper.

Acknowledgments

The authors are grateful to the National Counsel of Technological and Scientific Development (CNpQ) for the Grant (Process no. 481519/2013-0).

References

  1. L. Bjørndal, C. Reit, G. Bruun et al., “Treatment of deep caries lesions in adults: randomized clinical trials comparing stepwise vs. direct complete excavation, and direct pulp capping vs. partial pulpotomy,” European Journal of Oral Sciences, vol. 118, no. 3, pp. 290–297, 2010. View at Publisher · View at Google Scholar · View at Scopus
  2. L. Bjørndal, T. Larsen, and A. Thylstrup, “A clinical and microbiological study of deep carious lesions during stepwise excavation using long treatment intervals,” Caries Research, vol. 31, no. 6, pp. 411–417, 1997. View at Google Scholar · View at Scopus
  3. L. Bjørndal and A. Thylstrup, “A practice-based study on stepwise excavation of deep carious lesions in permanent teeth: a 1-year follow-up study,” Community Dentistry and Oral Epidemiology, vol. 26, no. 2, pp. 122–128, 1998. View at Google Scholar · View at Scopus
  4. E. A. M. Kidd, “How 'clean' must a cavity be before restoration?” Caries Research, vol. 38, no. 3, pp. 305–313, 2004. View at Publisher · View at Google Scholar · View at Scopus
  5. E. Leksell, K. Ridell, M. Cvek, and I. Mejàre, “Pulp exposure after stepwise versus direct complete excavation of deep carious lesions in young posterior permanent teeth,” Endodontics and Dental Traumatology, vol. 12, no. 4, pp. 192–196, 1996. View at Google Scholar · View at Scopus
  6. M. Maltz, L. S. Alves, J. J. Jardim, M. Dos Santos Moura, and E. F. De Oliveira, “Incomplete caries removal in deep lesions: a 10-year prospective study,” American Journal of Dentistry, vol. 24, no. 4, pp. 211–214, 2011. View at Google Scholar · View at Scopus
  7. M. Maltz, R. Garcia, J. J. Jardim et al., “Randomized trial of partial vs. stepwise caries removal: 3-year follow-up,” Journal of Dental Research, vol. 91, pp. 1026–1031, 2012. View at Google Scholar
  8. M. Maltz, E. F. Oliveira, V. Fontanella, and G. Carminatti, “Deep caries lesions after incomplete dentine caries removal: 40-month follow-up study,” Caries Research, vol. 41, no. 6, pp. 493–496, 2007. View at Publisher · View at Google Scholar · View at Scopus
  9. K. L. Weerheijm and H. J. Groen, “The residual caries dilemma,” Community Dentistry and Oral Epidemiology, vol. 27, no. 6, pp. 436–441, 1999. View at Google Scholar · View at Scopus
  10. J. S. Paddick, S. R. Brailsford, E. A. M. Kidd, and D. Beighton, “Phenotypic and genotypic selection of microbiota surviving under dental restorations,” Applied and Environmental Microbiology, vol. 71, no. 5, pp. 2467–2472, 2005. View at Publisher · View at Google Scholar · View at Scopus
  11. M. Maltz, E. F. De Oliveira, V. Fontanella, and R. Bianchi, “A clinical, microbiologic, and radiographic study of deep caries lesions after incomplete caries removal,” Quintessence International, vol. 33, no. 2, pp. 151–159, 2002. View at Google Scholar · View at Scopus
  12. M. Maltz, S. L. Henz, E. F. de Oliveira, and J. J. Jardim, “Conventional caries removal and sealed caries in permanent teeth: a microbiological evaluation,” Journal of Dentistry, vol. 40, pp. 776–782, 2012. View at Google Scholar
  13. A. S. Pinto, F. B. De Araújo, R. Franzon et al., “Clinical and microbiological effect of calcium hydroxide protection in indirect pulp capping in primary teeth,” American Journal of Dentistry, vol. 19, no. 6, pp. 382–386, 2006. View at Google Scholar · View at Scopus
  14. A. I. Orhan, F. T. Oz, B. Ozcelik, and K. Orhan, “A clinical and microbiological comparative study of deep carious lesion treatment in deciduous and young permanent molars,” Clinical Oral Investigations, vol. 12, no. 4, pp. 369–378, 2008. View at Publisher · View at Google Scholar · View at Scopus
  15. C. Duque, T. D. C. Negrini, N. T. Sacono, D. M. P. Spolidorio, C. A. De Souza Costa, and J. Hebling, “Clinical and microbiological performance of resin-modified glass-ionomer liners after incomplete dentine caries removal,” Clinical Oral Investigations, vol. 13, no. 4, pp. 465–471, 2009. View at Publisher · View at Google Scholar · View at Scopus
  16. E. C. O. Lula, V. Monteiro-Neto, C. M. C. Alves, and C. C. C. Ribeiro, “Microbiological analysis after complete or partial removal of carious dentin in primary teeth: a randomized clinical trial,” Caries Research, vol. 43, no. 5, pp. 354–358, 2009. View at Publisher · View at Google Scholar · View at Scopus
  17. D. S. Wambier, F. A. Dos Santos, A. C. Guedes-Pinto, R. G. Jaeger, and M. R. L. Simionato, “Ultrastructural and microbiological analysis of the dentin layers affected by caries lesions in primary molars treated by minimal intervention,” Pediatric Dentistry, vol. 29, no. 3, pp. 228–234, 2007. View at Google Scholar · View at Scopus
  18. P. D. Marsh, “Are dental diseases examples of ecological catastrophes?” Microbiology, vol. 149, no. 2, pp. 279–294, 2003. View at Publisher · View at Google Scholar · View at Scopus
  19. R. A. Arthur, A. A. del Bel Cury, R. O. Mattos-Graner et al., “Genotypic and phenotypic analysis of S. mutans isolated from dental biofilms formed in vivo under high cariogenic conditions,” Brazilian Dental Journal, vol. 22, no. 4, pp. 267–274, 2011. View at Publisher · View at Google Scholar · View at Scopus
  20. Y. Li and P. W. Caufield, “Arbitrarily primed polymerase chain reaction fingerprinting for the genotypic identification of mutans streptococci from humans,” Oral Microbiology and Immunology, vol. 13, no. 1, pp. 17–22, 1998. View at Google Scholar · View at Scopus
  21. F. L. Lembo, P. L. Longo, C. Ota-Tsuzuki, C. R. M. D. Rodrigues, and M. P. A. Mayer, “Genotypic and phenotypic analysis of Streptococcus mutans from different oral cavity sites of caries-free and caries-active children,” Oral Microbiology and Immunology, vol. 22, no. 5, pp. 313–319, 2007. View at Publisher · View at Google Scholar · View at Scopus
  22. R. O. Mattos-Graner, M. H. Napimoga, K. Fukushima, M. J. Duncan, and D. J. Smith, “Comparative analysis of Gtf isozyme production and diversity in isolates of Streptococcus mutans with different biofilm growth phenotypes,” Journal of Clinical Microbiology, vol. 42, no. 10, pp. 4586–4592, 2004. View at Publisher · View at Google Scholar · View at Scopus
  23. L. B. Firmino, “Estudo da viabilidade bacteriana em dentina cariada selada,” in Programa de Pós-graduação em Odontologia, UFRGS, Porto Alegre, Brasil, 2011. View at Google Scholar
  24. T. Do, S. C. Gilbert, D. Clark et al., “Generation of diversity in streptococcus mutans genes demonstrated by MLST,” PLoS ONE, vol. 5, no. 2, Article ID e9073, 2010. View at Publisher · View at Google Scholar · View at Scopus
  25. P. Baca, A. M. Castillo, M. J. Liébana, F. Castillo, A. Martín-Platero, and J. Liébana, “Horizontal transmission of Streptococcus mutans in schoolchildren,” Medicina Oral, Patología Oral y Cirugía Bucal, vol. 17, pp. e495–e500, 2012. View at Google Scholar
  26. S. C. Mitchell, J. D. Ruby, S. Moser et al., “Maternal transmission of mutans streptococci in severe-early childhood caries,” Pediatric Dentistry, vol. 31, no. 3, pp. 193–201, 2009. View at Google Scholar · View at Scopus
  27. M. H. Napimoga, R. U. Kamiya, R. T. Rosa et al., “Genotypic diversity and virulence traits of Streptococcus mutans in caries-free and caries-active individuals,” Journal of Medical Microbiology, vol. 53, no. 7, pp. 697–703, 2004. View at Google Scholar · View at Scopus
  28. L. Grönroos and S. Alaluusua, “Site-specific oral colonization of mutans streptococci detected by arbitrarily primed PCR fingerprinting,” Caries Research, vol. 34, no. 6, pp. 474–480, 2000. View at Google Scholar · View at Scopus
  29. M. Bönecker, C. Toi, and P. Cleaton-Jones, “Mutans streptococci and lactobacilli in carious dentine before and after Atraumatic Restorative Treatment,” Journal of Dentistry, vol. 31, no. 6, pp. 423–428, 2003. View at Publisher · View at Google Scholar · View at Scopus
  30. C. S. Toi, M. Bönecker, and P. E. Cleaton-Jones, “Mutans streptococci strains prevalence before and after cavity preparation during Atraumatic Restorative Treatment,” Oral Microbiology and Immunology, vol. 18, no. 3, pp. 160–164, 2003. View at Publisher · View at Google Scholar · View at Scopus
  31. M. Saarela, J. Hannula, J. Mättö, S. Asikainen, and S. Alaluusua, “Typing of mutans streptococci by arbitrarily primed polymerase chain reaction,” Archives of Oral Biology, vol. 41, no. 8-9, pp. 821–826, 1996. View at Publisher · View at Google Scholar · View at Scopus
  32. Y. Li, P. W. Caufield, I. Redmo Emanuelsson, and E. Thornqvist, “Differentiation of Streptococcus mutans and Streptococcus sobrinus via genotypic and phenotypic profiles from three different populations,” Oral Microbiology and Immunology, vol. 16, no. 1, pp. 16–23, 2001. View at Publisher · View at Google Scholar · View at Scopus
  33. S. Kneist, F. Schmidt, A. Callaway et al., “Diversity of Lactobacillus species in deep carious lesions of primary molars,” European Archives of Paediatric Dentistry, vol. 11, no. 4, pp. 181–186, 2010. View at Google Scholar · View at Scopus
  34. I.-M. Redmo Emanuelsson and E. Thornqvist, “Distribution of mutans streptococci in families: a longitudinal study,” Acta Odontologica Scandinavica, vol. 59, no. 2, pp. 93–98, 2001. View at Publisher · View at Google Scholar · View at Scopus
  35. I.-M. R. Emanuelsson, P. Carlsson, K. Hamberg, and D. Bratthall, “Tracing genotypes of mutans streptococci on tooth sites by random amplified polymorphic DNA (RAPD) analysis,” Oral Microbiology and Immunology, vol. 18, no. 1, pp. 24–29, 2003. View at Publisher · View at Google Scholar · View at Scopus
  36. S. Hojo, M. Komatsu, R. Okuda, N. Takahashi, and T. Yamada, “Acid profiles and pH of carious dentin in active and arrested lesions,” Journal of Dental Research, vol. 73, no. 12, pp. 1853–1857, 1994. View at Google Scholar · View at Scopus
  37. J. A. McGrady, W. G. Butcher, D. Beighton, and L. M. Switalski, “Specific and charge interactions mediate collagen recognition by oral lactobacilli,” Journal of Dental Research, vol. 74, no. 2, pp. 649–657, 1995. View at Google Scholar · View at Scopus
  38. N. Takahashi and B. Nyvad, “Caries ecology revisited: microbial dynamics and the caries process,” Caries Research, vol. 42, no. 6, pp. 409–418, 2008. View at Publisher · View at Google Scholar · View at Scopus