SciELO - Scientific Electronic Library Online

 
vol.30 issue1Cytotoxic effects of new MTA-based cement formulations on fibroblast-like MDPL-20 cellsDoes a toothpaste containing blue covarine have any effect on bleached teeth? An in vitro, randomized and blinded study author indexsubject indexarticles search
Home Pagealphabetic serial listing  

Services on Demand

Journal

Article

Indicators

Related links

Share


Brazilian Oral Research

Print version ISSN 1806-8324On-line version ISSN 1807-3107

Braz. oral res. vol.30 no.1 São Paulo  2016  Epub Mar 08, 2016

http://dx.doi.org/10.1590/1807-3107BOR-2016.vol30.0030 

Original Research

Antiseptics and microcosm biofilm formation on titanium surfaces

Georgia VERARDIa 

Maximiliano Sérgio CENCIb 

Tamires Timm MASKEb 

Bruna WEBBERc 

Luciana Ruschel dos SANTOSc 

aUniversidade de Passo Fundo – UPF, Faculdade de Odontologia, Programa de Pós-Graduação em Odontologia, Passo Fundo, RS, Brazil.

bUniversidade Federal de Pelotas – UFPel, Faculdade de Odontologia, Programa de Pós-Graduação em Odontologia, Pelotas,RS, Brazil.

cUniversidade de Passo Fundo – UPF, Faculdade de Agronomia e Medicina Veterinária, Programa de Pós-Graduação em Bioexperimentação, Passo Fundo, RS, Brazil.

Abstract

Oral rehabilitation with osseointegrated implants is a way to restore esthetics and masticatory function in edentulous patients, but bacterial colonization around the implants may lead to mucositis or peri-implantitis and consequent implant loss. Peri-implantitis is the main complication of oral rehabilitation with dental implants and, therefore, it is necessary to take into account the potential effects of antiseptics such as chlorhexidine (CHX), chloramine T (CHT), triclosan (TRI), and essential oils (EO) on bacterial adhesion and on biofilm formation. To assess the action of these substances, we used the microcosm technique, in which the oral environment and periodontal conditions are simulated in vitro on titanium discs with different surface treatments (smooth surface - SS, acid-etched smooth surface - AESS, sand-blasted surface - SBS, and sand-blasted and acid-etched surface - SBAES). Roughness measurements yielded the following results: SS: 0.47 µm, AESS: 0.43 µm, SB: 0.79 µm, and SBAES: 0.72 µm. There was statistical difference only between SBS and AESS. There was no statistical difference among antiseptic treatments. However, EO and CHT showed lower bacterial counts compared with the saline solution treatment (control group). Thus, the current gold standard (CHX) did not outperform CHT and EO, which were efficient in reducing the biofilm biomass compared with saline solution.

Key words: Peri-Implantitis; Biofilms; Titanium

Introduction

Dental implants offer an alternative so that patients’ rehabilitation and esthetic needs can be met, but failures have been reported and attributed to peri-implantitis.1,2 Described as an inflammatory disease that affects tissues in the vicinity of the osseointegrated implant, peri-implantitis leads to bone loss3 and subsequent implant loss; however, bacterial adhesion and biofilm accumulation are the major causes of this disease.4,5

The basic treatment for periodontitis and peri-implantitis consists of debridement of the affected surface (dental implant).6,7 Implant surfaces are highly microstructured and macrostructured so that they increase osseointegration, but rough surfaces facilitate microbial adhesion and formation of complex biofilms, hindering the debridement of implant surfaces.4,6,7 Additional therapy with antibiotics and antiseptics has been proposed for the removal of pathogenic biofilms and improvement of nonsurgical treatment outcomes.6,7,8,9

The healing potential of peri-implant defects after suppression of peri-implant microbial biofilm has been reported; nevertheless, treatment recommendations and the most appropriate chemical agent for decontamination of implants are yet unsatisfying.10,6,7,8

Gosau et al.11 investigated the effects of sodium hypochlorite, hydrogen peroxide, chlorhexidine digluconate, citric acid, essential oils, and triclosan on in vivo biofilms in healthy individuals, concluding that antiseptics had bactericidal effects on microbial adhesion, but interindividual variations hampered the interpretation of results. Moreover, the microbiota on teeth or implants under healthy conditions has qualitative and quantitative differences from those observed on sites affected by peri-implantitis or periodontitis.12

Filoche et al.13 used the microcosm technique to assess biomass and bacterial viability after oral antiseptic treatments, concluding that biomass was reduced by all tested solutions, but viability remained unchanged. Other authors have used this technique with patients’ saliva since it simulates the oral cavity and is therefore appropriate for investigations in the field of cariology and for tests with antiseptic substances.14,15,16

In the present study, we assessed the effect of four antiseptics against microcosm biofilm on titanium surfaces with different treatments.

Methodology

The study protocol (no. 135.946) was approved by the Research Ethics Committee ofUniversidade de Passo Fundo - UPF.

Titanium specimens. Sterile titanium specimens were donated by Titanium Fix (Titanium Fix, São José dos Campos, Brazil) in discs measuring 5.5 mm in diameter and 2 mm in thickness. The following types of surface were analyzed: smooth surface (SS), acid-etched smooth surface (AESS), sand-blasted surface (SBS), and sand-blasted and acid-etched surface (SBAES).

Surface roughness measurement. Surface roughness was measured by a SurrCode SE1200 profilometer (Kosakalab, Tokyo, Japan), calibrated with V 200, H 25 mm/λc and λc 0.25 mm, and average roughness (Ra) was considered to be that provided by the equipment.

Microcosm biofilm formation . For in vitro biofilm formation by the microcosm technique, we used the model described by van de Sande et al.17 Saliva was collected from a non-smoking volunteer with periodontal disease who had not taken antibiotics in the previous month, who had abstained from oral hygiene in the past 24 h, and who had fasted for 2 h prior to collection, being then processed. This patient signed a written informed consent form authorizing his participation in the study. 90 mL of saliva stimulated with paraffin (Parafilm®, Bemis Company, Oshkosh, USA) was collected into a sterile graded collector and transported to the laboratory under refrigeration, stored in a sterile vessel and homogenized in a vortex mixer.18 After collection, the patient was referred to free-of-charge periodontal treatment at the Dental School ofUniversidade Federal de Pelotas - UFPel.

Artificial saliva (DMM) preparation. The defined medium mucin (DMM) was obtained as proposed by Wong and Sissons,16 consisting of porcine gastric mucin (2.5 g/l), urea (1.0 mmol/l), salts in mmol/l (CaCl2: 1.0; MgCl2: 0.2; KH2PO4: 3.5; K2HPO4: 1.5; NaCl: 10.0; KCl: 15.0; NH4Cl: 2.0), 21 free amino acids, 17 vitamins, and growth factors. The medium contains amino acids for the protein/peptide equivalent (in mmol/l), whose concentrations are based on that of human saliva: alanine (1.95), arginine (1.30), asparagine (1.73), aspartic acid (1.52), cysteine (0.05), glutamic acid (5.41), glutamine (3.03), glycine (1.95), histidine (1.08), isoleucine (2.38), leucine (3.68), serine (3.46), threonine (1.08), tryptophan (0.43), tyrosine (2.17), valine (2.38), and casein (5.0 g/l).

Biofilm growth. The patient’s saliva was inoculated onto microplated titanium specimens using 400 µL per well and incubated in a microbiological incubator at 37°C for 1 hour. Thereafter, the saliva was aspirated from the well bottoms and 1.8 mL of artificial saliva was added to each microplate. The plates were incubated at 37°C for 24 h in anaerobic jars (Anaerobac® - Probac do Brasil Produtos Bacteriológicos Ltda., São Paulo, Brazil) and biofilms were formed independently on titanium specimens. After that, the specimens were transferred with sterile tweezers to a new plate containing DMM and kept under anaerobic conditions (Anaerobac® jar) and allowed to rest for 24 hour.

Treatment with antiseptic substances. After 48 h of incubation, the specimens were transferred with sterile tweezers to a new plate containing 2 mL of each antiseptic and maintained in contact for 60 seconds. The tested antiseptics were: chlorhexidine 0.12% (Periogard - Colgate-Palmolive Company, São Paulo, Brazil), chloramine T (Trihydral - Perland Pharmacos Ltda., Londrina, Brazil), triclosan (Plax - Colgate-Palmolive Company, São Paulo, Brazil), and essential oils containing eucalyptol, thymol, methyl salicylate, and menthol (Listerine - Johnson & Johnson do Brasil Ind. e Com. de Produtos para Saúde Ltda., São Paulo, Brazil). Saline solution 0.9% (Laboratório Arboreto, Juiz de Fora, Brazil) was used for the control group.

Quantification of viable cells. After treatment with antiseptic substances, the specimens were removed from the wells with sterile tweezers, and non-adherent cells were washed off with 2 mL of sterile saline solution in microplates. The specimens, following the order of treatment of surface and of antiseptic, were placed in microtubes containing 1 mL of saline solution, stored on ice, homogenized in a vortex mixer (Phoenix Modelo AP 56 - Phoenix Indústria e Comércio de Equipamentos Científicos Ltda, Araraquara, Brazil) and sonicated (Sonicador Vibra Cell - Sonics and Materials, Danbury, USA) at 30 W using three pulses of 10 s at a 5-second interval in order to obtain a homogenous biofilm suspension.19

The suspensions were diluted in saline solution up to 10-7 and inoculated in duplicate in blood agar for the count of total microorganisms under anaerobic conditions (Anaerobac®) at 37°C for 96 hours. Colony-forming units were counted and the results were expressed in CFU/mm2.19

Statistical analysis. The CFU data were analyzed using two-way ANOVA at a 5% significance level and the data between the groups were compared by Tukey’s test. The software used was SigmaPlot Version 11.0, from Systat Software Inc., San Jose, USA.

Results

The following surface roughness values were obtained: 0.47 µm for SS, 0.43 µm for AESS, 0.79 µm for SBS, and 0.72 µm for SBAES.

There was a significant difference concerning antiseptic agents (p = 0.003) and titanium surfaces (p = 0.015), but the interaction between these factors was not statistically significant (p = 0.718). After 48 h of biofilm growth, a thick biomass was observed on the titanium surfaces. Bacterial counts were higher on SBS, on SBAES, and on SS than on AESS. There was statistical difference only between SBS and AESS (Table).

Table Average bacterial counts (SE) on titanium surfaces submitted to different treatments. 

Surface CFU/ mm2 (log)
Sand-blasted (SBS) 8.16 (0.09)a
Sand-blasted and acid-etched (SBAES) 8.05 (0.87)a
Smooth (SS) 8.01 (0.86)ab
Acid-etched smooth (AESS) 7.97 (0.86)b

Values in the column bearing the same letter are statistically similar (p > 0.05).

There was no statistical difference among antiseptics in the groups treated with triclosan (TRI), chlorhexidine (CHX), essential oils (EO), and chloramine T (CHT). However, there were statistical differences between EO and CHT when compared with saline solution (Figure).

Note: AESS: acid-etched smooth surface; SBAES: sand-blasted and acid-etched surface; SBS: sand-blasted surface; SS: smooth surface. Considering the treatment performed, the asterisks show statistical difference between titanium surfaces. Considering titanium surfaces, the brackets represent statistical difference between antiseptic treatments (p < 0.05).

Figure Average bacterial counts (SE) considering antiseptic treatments and titanium surfaces. 

Discussion

Berglundh et al.20 investigated SBAES in spontaneous progression of peri-implantitis in dogs and concluded that progression of peri-implantitis is more pronounced on rough surfaces than on smooth ones. In this context, Pongnarisorn et al.21 assessed the effects of different surface treatments on the transmucosal area of implants (machined, acid-etched, and anodized) and suggested that the development of implant-associated inflammation is not dependent upon the type of surface or roughness, but rather upon the presence of bacterial plaque. The authors also mention that the type of surface does not interfere with the quality of the inflammatory infiltrate, with predominance of T cells in all cases, and does not interfere with the microbiota around the implant, although the presence of notches in the subgingival area predisposes to plaque accumulation, thereby increasing the inflammatory infiltrate.

Chlorhexidine was not so good as the other substances used. For over three decades, CHX had been the gold standard, compared with other chemical agents, because it prevented the formation of dental biofilm;22 however, the present study demonstrates that it is less efficient in reducing microbial colonization. One of the advantages of its use is its broad spectrum and its prolonged and continuous substantivity, even in the presence of blood and other bodily fluids.23 It produces adverse effects, but no systemic toxicity has been reported so far,24 as it is poorly absorbed by the gastrointestinal tract. Nevertheless, its prolonged used may cause temporary clinical side effects, such as extrinsic staining of teeth, of restorations, and of the tongue, desquamation of the oral mucosa and, occasionally, allergic reactions.

Hanke,25 Sweet et al.,26 and Rams et al.7 comment that CHT significantly reduces bacterial colonization, especially in peri-implantitis and in post-extraction bacteremia. In the present study, CHT was found to have the same microbial growth reduction as that provided by CHX and was effective when compared with the control group (Figure). The group treated with EO had fewer bacterial counts than those treated with saline solution, which is in line with the results obtained by Bugno et al.,27 who found that the antimicrobial and antifungal activity of essential oils was better than that of CHX. However, Monfrin and Ribeiro28 and Moreira et al.29 observed that Listerine was less efficient in reducing the microbiota in the saliva, which is inconsistent, who mention that essential oils in long-term clinical trials were efficient and safe.

Even though there was no statistically significant difference between the assessed antiseptics, when an implantologist recommends a mouthwash solution, he/she may choose one that is good at reducing bacterial count and has no adverse effects in the long run. There appears to be consensus agreement that the use of prophylactic antiseptics should be a complement rather than a substitute for conventional mechanical methods, thus adding to and trying to eliminate the deficiencies of mechanical oral hygiene habits.6

Conclusion

Different antiseptics reduce the amount of bacteria in titanium implants, but CHX, the current gold standard, was not as good against microcosm biofilm formation as CHT and EO, which were efficient in reducing the biofilm biomass compared with saline solution.

Acknowledgments

We thank Titanium Fix for providing the titanium specimens used in this study andFundação de Amparo à Pesquisa do Estado do Rio Grande do Sul - FAPERGS PqG 12/2523-5 for the partial financial support.

References

1. Lindhe J, Meyle J. Peri-implant diseases: consensus report of the sixth European workshop on periodontology; Group D of European Workshop on Periodontology. J Clin Periodontol. 2008;35(Suppl 8):282-5. doi:10.1111/j.1600-051X.2008.01283.x [ Links ]

2. Lopes AC, Rezende CEE, Fernandes MS, Weinfeld I. Bacterial leakage of the implant-abutment interface: what the implantologist should know. Rev Gaucha Odontol. 2010;58(2);239-42. Portuguese. [ Links ]

3. Heuer W, Elter C, Demling A, Neumann A, Suerbaum S, Hannig M, et al. Analysis of early biofilm formation on oral implants in man. J Oral Rehabil. 2007;34(5):377-82. doi:10.1111/j.1365-2842.2007.01725.x [ Links ]

4. Bürgers R, Gerlach T, Hahnel S, Schwarz F, Handel G, Gosau M. In vivo and in vitro biofilm formation on two different titanium implant surfaces. Clin Oral Implants Res. 2010;21(2);156-64. doi:10.1111/j.1600-0501.2009.01815.x [ Links ]

5. Lee BC, Jung GY, Kim DJ, Han JS. Initial bacterial adhesion on resin, titanium and zirconia in vitro. J Adv Prosthodont. 2011;3(2)81-4. doi:10.4047/jap.2011.3.2.81 [ Links ]

6. Lindhe J, Lang NP, Karring T, editors. Tratado de Periodontia Clínica e Implantodontia Oral. Rio de Janeiro: Guanabara Koogan; 2005. [ Links ]

7. Rams TE, Keyes PH, Jenson AB. Morphological effects of inorganic salts, chloramine-T, and citric acid on subgingival plaque bacteria. Quintessence Int Dent Dig. 1984;15(8):835-44. [ Links ]

8. Renvert S, Roos-Jansaker AM, Claffey N. Non-surgical treatment of peri-implant mucositis and peri-implantitis: a literature review. J Clin Periodontol. 2008;35(Suppl 8):305-15. doi:10.1111/j.1600-051X.2008.01276.x [ Links ]

9. Rosenthal S, Spangberg L, Safavi K. Chlorhexidine substantivity in root canal dentin. Oral Surg Oral Med Oral Pathol Oral Radiol Endod. 2004;98(4):488-92. [ Links ]

10. Klinge B, Gustafsson A, Berglundh T. A systematic review of the effect of anti-infective therapy in the treatment of periimplantitis. J Clin Periodontol. 2002;29(Suppl 3):213-25. doi:10.1034/j.1600-051X.29.s3.13.x [ Links ]

11. Gosau M, Hanhel S, Schwarz F, Gerlach T, Reichert TE, Bürgers R. Effect of six different peri-implantitis disinfection methods on in vivo human oral biofilm. Clin Oral Implants Res. 2010;21(8):866-72. doi:10.1111/j.1600-0501.2009.01908.x [ Links ]

12. Leonhardt A, Bergstrom C, Lekholm U. Microbiologic diagnostics at titanium implants. Clin Implant Dent Relat Res. 2003;5(4):226-32. doi:10.1111/j.1708-8208.2003.tb00205.x [ Links ]

13. Filoche SK, Coleman MJ, Angker L, Sissons CH. A fluorescence assay to determine the viable biomass of microcosm dental plaque biofilms. J Microbiol Methods. 2007;69(3)489-96. doi:10.1016/j.mimet.2007.02.015 [ Links ]

14. Filoche SK, Soma D, Van Bekkum M, Sissons CH. Plaques from different individuals yield different microbiota responses to oral-antiseptic treatment. FEMS Immunol Med Microbiol. 2008;54(1):27-36. doi:10.1111/j.1574-695X.2008.00443.x [ Links ]

15. Schwarz F, Bieling K, Bonsmann M, Latz T, Becker J. Nonsurgical treatment of moderate and advanced periimplantitis lesions: a controlled clinical study. Clin Oral Investig. 2006;10(4):279-88. doi:10.1007/s00784-006-0070-3 [ Links ]

16. Wong L, Sissons CH. A comparison of human dental plaque microcosm biofilms grown in an undefined medium and chemically defined artificial saliva. Arch Oral Biol. 2001;46(6):477-86. doi:10.1016/S0003-9969(01)00016-4 [ Links ]

17. van de Sande FH, Azevedo MS, Lund RG, Huysmans MCDNJM, Cenci MS. An in vitro biofilm model for enamel demineralization and antimicrobial dose-response studies. Biofouling. 2011;27(9):1057-63. doi:10.1080/08927014.2011.625473 [ Links ]

18. Filoche SK, Soma KJ, Sissons CH. Caries-related plaque microcosm biofilms developed in microplates. Oral Microbiol Immunol. 2007;22(3):73-9. doi:10.1111/j.1399-302X.2007.00323.x [ Links ]

19. Darouiche RO. Treatment of infections associated with surgical implants. N Engl J Med. 2004;350(14):1422-29. doi:10.1056/NEJMra035415 [ Links ]

20. Berglundh T, Gotfredsen K, Zitzmann NU, Lang NP, Lindhe J. Spontaneous progression of ligature induced peri-implantitis at implants with different surface roughness: an experimental study in dogs. Clin Oral Implants Res. 2007;18(5):655-61. doi:10.1111/j.1600-0501.2007.01397.x [ Links ]

21. Pongnarisorn NJ, Gemmell E, Tan AE, Henry PJ, Marshall RI, Seymour GJ. Inflammation associated with implants with different surface types. Clin Oral Implants Res. 2007;18(1):114-25. [ Links ]

22. Charles CH, Mostler KM, Bartels LL, Mankodi SM. Comparative antiplaque and antigingivitis effectiveness of a chlorhexidine and an essential oil mouthrinse: 6-month clinical trial. J Clin Periodontol. 2004;31(10):878-84. doi:10.1111/j.1600-051X.2004.00578.x [ Links ]

23. Roos-Jansaker AM, Renvert S, Egelberg J. Treatment of periimplant infections: a literature review. J Clin Periodontol. 2003;30(6):467-85. doi:10.1034/j.1600-051X.2003.00296.x [ Links ]

24. Ciancio SG. Chemical agents: plaque control, calculus reduction and treatment of dentinal hypersensitivity. Periodontol 2000. 1995;8(1):75-86. doi:10.1111/j.1600-0757.1995.tb00046.x [ Links ]

25. Hanke MT. Studies on the local factors in dental caries. I. Destruction of plaques and retardation of bacterial growth in the oral cavity. J Am Dent Assoc. 1940;27(9):1379-93. doi:10.14219/jada.archive.1940.0269 [ Links ]

26. Sweet JB, Gill VJ, Chusid MJ, Elin RJ. Nitroblue tetrazolium and limulus assays for bacteremia after dental extraction: effect of tropical antiseptics. . J Am Dent Assoc. 1978;96(2):276-81. [ Links ]

27. Bugno A, Nicolleti MA, Almodóvar AAB, Pereira TC, Auricchio MT. Enxaguatórios bucais: avaliação da eficácia antimicrobiana de produtos comercialmente disponíveis. Rev Inst Adolfo Lutz. 2006;65(1):40-5. [ Links ]

28. Monfrin RCP, Ribeiro MC. Avaliação in vitro de anti-sépticos bucais sobre a microbiota da saliva. Rev Assoc Paul Cir Dent. 2000;54(5):401-7. [ Links ]

29. Moreira ACA, Pereira MHQ, Porto MR, Rocha LAP, Nascimento BC, Andrade P M. Avaliação in vitro da atividade antimicrobiana de antissépticos bucais. Rev Cienc Med Biol. 2009;8(2):153-61. [ Links ]

Received: May 26, 2015; Revised: September 21, 2015; Accepted: November 16, 2015

Corresponding Author: Luciana Ruschel dos Santos. E-mail:luruschel@upf.br

Declaration of Interests: The authors certify that they have no commercial or associative interest that represents a conflict of interest in connection with the manuscript.

Creative Commons License This is an Open Access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.