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Brazilian Journal of Oral Sciences, Vol. 8, No. 1, Jan-Mar, 2009, pp. 2-8 Genetic polymorphism of Streptococcus mutans strains associated with incomplete caries removal Cristiane Duque1, Thais de Cássia Negrini2, Nancy Tomoko Sacono3, Marcelo Fabiano Gomes Boriollo4, José Francisco Hofling5, Josimeri Hebling6, Denise Madalena Palomari Spolidorio7 1 DDS, PhD, Post-doctorate student, Department of Microbiology and Immunology, Faculdade de Odontologia de Piracicaba, Universidade Estadual de Campinas (Unicamp), Piracicaba (SP), Brazil; Department of Orthodontics and Pediatric Dentistry, Faculdade de Odontologia de Araraquara, Universidade Estadual Paulista “Júlio de Mesquita Filho” (Unesp), Araraquara (SP), Brazil
Received for publication: March 15, 2009 Code Number: os09001 Abstract Aim: Despite the antibacterial properties of dental materials, the survival of residual bacteria under restorations has been demonstrated after incomplete caries removal. The aim of this study was to evaluate the genetic polymorphism of Streptococcus mutans strains isolated from deep dentinal lesions before and three months after incomplete caries removal. Keywords: Streptococcus mutans, polymerase chain reaction, polymorphism, genetic, dental caries. Introduction The wide distribution and variety of oral bacteria demonstrate their ability to survive among their human hosts, as a result of the efficient transmission of strains and their persistence in the oral cavity1. There are several microenvironments within the mouth that harbor communities of pathogenic or non-pathogenic bacteria, called biofilms, which are mainly found in the hard surfaces of the teeth2. The biochemical instability between tooth substance and the overlying biofilm, determined by intense production of acids derived from bacterial metabolism, and may lead to dissolution of the dental hard tissues and consequently to the development of a carious lesion3. Among the oral pathogens, mutans streptococci (MS) comprise a group of seven bacterial species of which only Streptococcus mutans, S. sobrinus, S. rattus and S. cricetus can be found on the human oral cavity4. The highest prevalence of S. mutans (74 to 94%) among oral streptococci, isolated from carious lesions demonstrates their association with caries development5. S. mutans has the ability of tolerating continuous cycles of acid shock, produced in the oral environment by mechanisms of proton extrusion from the cell via membrane-associated processes and by acid end product efflux, protecting important cellular components, especially DNA, from aggressive effects of acidification6. In addition, in adverse environments with nutrient restriction, some bacteria, such as S. mutans, are able to obtain carbohydrates from host-derived saliva or serum glycoproteins7-8. Genetic diversity among strains of the same species may reflect their ability to survive extreme environmental changes. Therefore, selection of strains best fitted to a given environment can be related to the generation of genetic variants, representing clonal populations, which conserve DNA into single cell lines, or non-clonal populations, which incorporate DNA from other cells9. Several studies have shown genetic heterogeneity, among S. mutans strains, using modern molecular typing techniques, such as arbitrary primed polymerase chain reaction (AP-PCR), multilocus enzyme electrophoresis (MLEE), restriction fragment length polymorphisms (RFLP) and repetitive extragenic palindromic PCR (REP-PCR)9-15. AP-PCR is a valuable DNA analysis tool that have been used mainly in streptococcal epidemiology and transmission studies, by virtue of its rapidity, efficiency and reproducibility in generating genetic fingerprints of bacterial isolates10,16. Several studies using AP-PCR have demonstrated that caries-active children have greater genotypic diversity of S. mutans, compared to caries-free children14,17. In addition, a site-specific colonization pattern of S. mutans genotypes in coronal and root caries lesions has been reported12,18. The survival of oral streptococci has been observed even months or years after incomplete caries removal and cavity sealing19-21, suggesting the generation of treatment-resistant strains. Paddick et al.15 have detected different genotypes of Streptococcus oralis and Actinomyces naeslundii, five months after incomplete carious dentin removal, with reduced phenotypic and genotypic diversity compared to strains isolated from the initial samples before treatment. Therefore, the aim of this study was to evaluate the genetic polymorphism of S. mutans strains isolated from deep dentinal lesions before and three months after incomplete caries removal. Materials and methods Thirty-three molars (27 primary and 6 young permanent) with deep occlusal caries lesions were obtained from 20 children of both genders, aged four to ten years and subjected to indirect pulp treatment. The absence of irreversible pulpal or periapical diseases was determined through clinical and radiographic examinations of all selected teeth. This study was conducted under a protocol approved by the Research Ethics Committee of Faculdade de Odontologia de Araraquara of Universidade Estadual Paulista, in Brazil, “Júlio de Mesquita Filho” (Unesp) and a signed informed consent was obtained from the legal tutors for the the children’s participation. Clinical procedures and microbiological sampling Indirect pulp treatment was divided in two sessions. At the first session, anesthesia was delivered and a rubber dam applied to isolate the tooth and prevent contamination by saliva. Rubber cup pumice prophylaxis and antisepsis of operative area, using 0.2% chlorhexidine digluconate, were performed. If necessary, access to infected dentin was gained using a sterile # 245 carbide bur (KG Sorensen, Barueri, SP, Brazil) at high speed. After removing the superficial necrotic dentin, an initial collect (A) of carious dentin was sampled with a sterile excavator, immediately immersed in 1 mL of saline and maintained in refrigerated boxes (4 ºC). A sterile round steel bur at low speed was used to clean all carious tissue from the dentinoenamel junction, leaving a layer of soft dentin on the cavity floor to avoid pulp exposure. After washing and air-drying the cavities to remove debris, a second collect (B) of carious dentin was obtained from the mesial portion of the cavity floor for microbiological analysis, as previously described. In order to standardize the amount of collected dentin, a small cavity was created at the flat end of an amalgam plugger, which was completely filled with dentin. The remaining carious dentin was covered with resin-modified glass ionomer cements (Vitrebond; 3M/ESPE, St. Paul, MN, USA or Fuji Lining LC; GC Corp., Tokyo, Japan) or a calcium hydroxide-based cement (Dycal; Dentsply, Milford, DE, USA). The cavities were restored with temporary zinc oxide-eugenol cement (IRM; Dentsply). After three months, the second session of the indirect pulp treatment was undertaken. Under the same initial conditions of anesthesia and isolation, teeth were reopened and the liner materials, carefully and completely removed. A third collect (C) of carious dentin was sampled from the distal portion of the cavity floor, as already described. Later, the teeth were lined with glass ionomer cement and restored definitively with silver amalgam (Dispersalloy, Dentsply International, USA). Microbiological procedures The tubes containing the carious dentin samples were vortexed for two minutes and the suspension was serially diluted with saline. For each dilution, 25 μL of the samples were placed duplicated in agar sacarose bacitracin (SB) – SB-2022 and incubated at 37 oC for 24 hours. After incubation, six to eight representative colonies of streptococci were collected and inoculated individually in 5 mL of brain heart infusion (BHI) (Brain Heart Infusion, Difco Laboratories, BD, Sparks, MD, USA) broth for 24 hours. The purity of the cultures were confirmed using Gram technique and aliquots of the subculture frozen at -20 oC, in 10% glycerol BHI for posterior molecular analysis of bacterial isolates. Polymerase chain reaction The extraction of chromosomal DNA was described by Nociti et al.23 and modified by Nascimento et al.18. Briefly, overnight cultures in BHI broth were centrifuged, followed by washing twice with TRIS-EDTA buffer (10 mM Tris-HCl, 1 mM EDTA, pH = 8.0) and boiled for ten minutes. After centrifugation, 60 μL of supernatant was collected and used as templates for the PCR. In order to confirm S. mutans molecular identity, DNA from MS isolates obtained from carious dentin collects was submitted to PCR method, using specific primers for portions of the glucosyltransferase B gene (gtfB) following the bases sequences’: 5’ – ACT ACA CTT TCG GGT GGC TTG G – 3’ e 5’ – CAG TAT AAG CGC CAG TTT CAT C – 3’, to amplify a517 bp DNA fragment24. Each PCR mixture contained 5 μL of the DNA template, 5 µL of 10x PCR amplification buffer (100 mM Tris-HCl, 500 mM KCl, pH = 8.3), 0.2 mM of dNTPs (DNA polymerization mix), 3.0 mM MgCl2, 1 μM of each primer, 2.5 U of Taq DNA polymerase and sterile distilled water, in order to make a final volume of 25 µL. All PCR reagents were obtained from Invitrogen, Life Technologies, São Paulo, Brazil. Positive and negative controls of PCR were purified in genomic DNA of S. mutans (ATCC 25175) and sterile water, respectively. The amplification of DNA was performed in a thermocycler (GeneAmp PCR System 2400, Perkin-Elmer’s Applied Biosystems Division, Foster City, CA, USA) with initial denaturation at 95 oC for five minutes, followed by 30 cycles of denaturation at 95o C for 30 seconds, annealing at 59 oC for 30 seconds, extension at 72 oC for one minute, ending with final extension at 72 oC for seven minutes24. The PCR amplification products were separated by electrophoresis in 1% agarose gels in Tris-borato-EDTA (TBE), running buffer (pH = 8.0) at 75 V for two hours. Gels were stained with 0.5 μg of ethidium bromide/mL and visualized under ultraviolet light illumination (UltraLum; Labtrade do Brasil, São Paulo, Brazil). A 100 bp DNA ladder was included as a molecular-size marker in each gel. AP-PCR Strains identified as S. mutans by PCR method were used for genotyping. AP-PCR amplification was performed with primer OPA-13 (5’- CAGCACCCAC – 3’)10,14,18. All reactions were processed in a volume of 25 µL, containing 2.5 μL of 10x PCR amplification buffer (100 mM Tris-HCl, 500 mM KCl, pH = 8.3), 7 mM MgCl2, 0.2 μM of dNTP, 1 μM of primer, 2.5 U of Taq DNA polymerase, 50 ηg of DNA and distilled water18. The amplification was performed in the same thermocycler with an initial denaturation at 94 oC for five minutes and 45 cycles of denaturation at 94 oC for 30 seconds, annealing at 36 oC for 30 seconds, extension at 72 oC for one minute, ending with final extension at 72 oC for three minutes. Amplicons generated by AP-PCR were analyzed electrophoretically in 1.4% agarose gel, in TBE running buffer and stained in 0.5 µg/mL ethidium bromide. A 1 Kb DNA ladder was used as molecular-size marker. The gels were photographed and their images, captured with a digital imaging system (Kodak Digital Science 1D; Eastman Kodak Company, Rochester, NY, USA). The molecular weights for each band or amplicon were computed and analyzed using the Sigma Gel software (Jandel Scientific, San Rafael, CA, USA). The amplicons were converted in binary data and submitted to NTSYS-pc software (Applied Biostatistics, Inc., Setauket, NY, USA), using the simple matching coefficient (SSM), and the unweighted pair group method with mathematic average (UPGMA) cluster analysis to generate similarity dendrograms. Results Only ten deciduous teeth were included in the study that showed positive culture in all collects (A, B and C). From the 429 selected MS colonies, 377 (87.9%) were identified as S. mutans. Table 1 shows the number of MS isolates and S. mutans strains, according to the collected period and material group. One hundred and seventy-three S. mutans isolates were chosen for AP-PCR analysis considering the three collects (A, B and C). Because the distribution of the teeth was not similar among the material groups, data analysis was performed for all teeth, independent of the lining material. All of them were subjected to AP-PCR. Each isolate received a specific code according to the following sequence: number of the tooth (from 1 to 10), collect (A, B and C) and number of the bacterial isolate (from one to eight), for example: 5A2. The amplification of genomic DNA of this species resulted in fragments (amplicons or eletrophoretic bands), ranging from 0.3 to 2.2 Kb in size. Some of them were species-specific, whereas others were found in only one or few S. mutans strains. The AP-PCR fingerprinting profile analysis with primer OPA-13 showed distinct genotypes patterns of S. mutans obtained from caries samples. In this study, the same genotypic pattern was considered for identical or highly related samples with genetic similarity (SSM) ≥ 0.869 (threshold). Considering each tooth separately, a maximum of nine genotypic patterns were found in each tooth from all the collects (A, B and C). In addition, at least one genotypic pattern was repeated in the three collects. Table 2 shows the total number of genotypic patterns detected in each tooth, without repetitions among the collects (genetic similarity SSM ≥ 0.869). Based in the matrices generated by the UPGMA analysis using coefficient SSM, the genetic similarity levels obtained among the S. mutans strains are illustrated in Figure 1, which shows representative dendrograms of S. mutans isolates obtained for each tooth (1 to 10). The genetic similarity values ranged from 0.555 ≤ SSM ≤ 1. Two to four S. mutans groups (clusters) containing identical or highly related isolates (SSM ≥ 0.869) were found in each tooth. These groups can be identified by tonalities of gray in the dendrograms. Identical or highly related isolates were found in all collects in the same tooth. Comparing the genotypic patterns in Collect C, which were obtained three months after dental treatment, a dendrogram with the SM isolates is shown in Figure 2 and the genetic similarity index observed was 0.621 ≤ SSM ≤ 1. UPGMA analysis revealed 11 groups containing identical or highly related isolates (0.869 ≤ SSM ≤ 1). Seven of these groups had isolates corresponding to a specific tooth (Groups 1, 3, 7, 8, 9, 10 and 11). Other three groups had isolates from two distinct teeth (Groups 4, 5 and 6) and one group had isolates from four teeth (Group 2), demonstrating genetic similarity among S. mutans strains, obtained from the same tooth and greater polymorphism among isolates from different teeth. Discussion S. mutans is the major pathogen associated with dental caries in humans. Several studies have demonstrated its capacity to tolerate extreme pH changes, varying from alkalinity25 to high levels of acidity6. Furthermore, in situations of environmental nutrient stress, such as after cavity sealing, S. mutans is capable of producing glycosidic enzymes that release carbohydrates from serum glycoproteins7-8 present in the dentinal tubules15. Therefore, it is plausible to suggest that specific phenotypic characteristics could be expressed by genotypes best fitted to survive and/or grow in adverse environments1. In the present study, three months after incomplete caries removal, genetic diversity of S. mutans strains was observed in samples of deep caries lesions even after the contact with antibacterial materials, such as glass-ionomer cements26. Analyzing each tooth individually, a maximum of nine different genotypic patterns was observed from all the collects. Several studies have demonstrated genetic heterogeneity among S. mutans strains, obtained from saliva or dental biofilm samples10,13, especially in caries-active individuals14,17. Some investigators have suggested that the genotype frequency can even vary among the oral sites, but a site-specific colonization of genotypes seems to exist in dental caries lesions12,18. Some identical or highly genotypic patterns were isolated in more than one collect, in a same tooth. Although few clones (SSM = 1) have been detected, the great genetic similarity could indicate resistance of S. mutans strains. Genetic groups containing highly related S. mutans isolates are thought to have derived from a single ancestral cell9, which could have originated new strains with genetic variations. It is commonly accepted that genetic polymorphism between close species is determined by modifications in base pairing, by deletion or insertion of new genetic sequences27, in addition to clone transmission to external sources28. However, the frequency of these events in vivo is not known yet8. As a part of the evolution process, bacteria have the ability to gain genetic material from other cells using the mechanism of transformation. For recombination to occur, the foreign DNA must share between 70 and 100% identity with the sequence in the recipient strain’s chromosome29. Some S. mutans strains may acquire several cariogenic properties30, fluoride31 and antibiotic resistance32 by transformation. It is possible that S. mutans could act as a donor of DNA to another species, such as S. sanguis and S. milleri33. Cvitkovitch2 suggested that bacterial transformations can occur in environments which experience extreme changes and fluctuations in population dynamics, such as the oral cavity. Li et al.34 have demonstrated that S. mutans cells are hypertransformable when grown in biofilms in vitro. Bacteria in these environments are frequently exposed to various stress conditions, such as nutrient excess or shortage, low pH, high osmolarity and the consumption of antimicrobial agents by the host9,34. Therefore, natural genetic transformation could be considered an important mechanism of cell’s adaptation to environmental changes, providing microbial resistance, genetic variation and rapid evolution of the virulence factors2,29,34. Comparing S. mutans genotypic patterns isolated from carious samples after three months of the dental treatment, genetic diversity was observed among the teeth and high intra-tooth similarity was detected. These results may suggest that different resistant S. mutans strains have tooth-specific colonization, because it was not possible to verify a common genetic pattern within the sample subjected to the treatment proposed in this study. Some investigators have shown genetic similarity among S. mutans genotypes obtained from members of the same family11,13,35-36 or individuals that cohabit in environments, such as nursery schools, denoting horizontal transmission37. Generally, non-related subjects rarely share identical S. mutans genotypes10. Nascimento et al.18 have compared 40 genetic types determined by AP-PCR, isolated from nine different patients and it was observed a great diversity among them, demonstrating that the maximum value of the similarity indices (SSM = 0.960) was observed only in isolates from the same individual. Despite some deficiencies, such as the difficulty to visualize low-intensity bands and the need for more than one primer to increase the technique accuracy14, studies have shown the efficacy of AP-PCR in the detection of genetic polymorphism of various bacterial species10,16, obtaining similar results to more sophisticated techniques like MLEE14. In this study, genetic heterogeneity was verified among S. mutans strains isolated form caries lesions, even three months after incomplete caries removal. The persistence of some identical genotypes and high genetic similarity among isolates of the same tooth, in distinct collects, denoted resistance of some S. mutans strains to dental treatment. The polymorphism observed in different teeth may suggest that resistant strains are specific to each tooth, since a common genetic pattern among individuals was not found. However, further studies are necessary to evaluate the phenotypic characteristics of different S. mutans genotypes resistant to indirect pulp treatment or another incomplete caries removal technique, observing possible similarities in the production of enzymes that participate of several virulence mechanisms of this species. Acknowledgements The authors thank Andréia Cristina Celi and Carina Bento Luis Macera for laboratorial assistance. This work was supported by the Brazilian agencies: Fundação de Amparo à Pesquisa do Estado de São Paulo (Fapesp) by grant 04/00677-1, and Coordenação de Aperfeioçoamento de Pessoal de Nível Superior (Capes). References
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