Accepted Manuscript Title: Influence of a Brazilian wild green propolis on the enamel mineral loss and Streptococcus mutans’ count in dental biofilm Author: Julia Gabiroboertz Cardoso Natalia Lopes Pontes Iorio Lu´ıs Fernando Rodrigues Maria Luiza Barra Couri Adriana Farah Lucianne Cople Maia Andr´ea Gonc¸alves Antonio PII: DOI: Reference:
S0003-9969(16)30028-0 http://dx.doi.org/doi:10.1016/j.archoralbio.2016.02.001 AOB 3544
To appear in:
Archives of Oral Biology
Received date: Revised date: Accepted date:
29-5-2015 19-12-2015 1-2-2016
Please cite this article as: Cardoso Julia Gabiroboertz, Iorio Natalia Lopes Pontes, Rodrigues Lu´is Fernando, Couri Maria Luiza Barra, Farah Adriana, Maia Lucianne Cople, Antonio Andr´ea Gonc¸alves.Influence of a Brazilian wild green propolis on the enamel mineral loss and Streptococcus mutans’ count in dental biofilm.Archives of Oral Biology http://dx.doi.org/10.1016/j.archoralbio.2016.02.001 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
1 Influence of a Brazilian wild green propolis on the enamel mineral loss and Streptococcus mutans’ count in dental biofilm Julia Gabiroboertz Cardoso1, Natalia Lopes Pontes Iorio2, Luís Fernando Rodrigues3, Maria Luiza Barra Couri4, Adriana Farah5, Lucianne Cople Maia6, Andréa Gonçalves Antonio7 1 Graduate
student, Department of Pediatric Dentistry and Orthodontics, Dental School,
Federal University of Rio de Janeiro, RJ, Brazil. 2
PhD, Adjunct Professor, Department of Basic Science, Federal Fluminense
University, Nova Friburgo Campus, Brazil. 3
Graduate student, Department of Basic Science, Federal Fluminense University,
Nova Friburgo Campus, Brazil. 4
Graduate student, Department of Basic Science, Federal Fluminense University,
Nova Friburgo Campus, Brazil. 5 PhD,
Adjunct Professor, Institute of Nutrition, Federal University of Rio de Janeiro, RJ,
Brazil. 6
PhD, Full Professor, Department of Pediatric Dentistry and Orthodontics, Dental
School, Federal University of Rio de Janeiro, RJ, Brazil. 7
PhD, Adjunct Professor, Department of Pediatric Dentistry and Orthodontics, Dental
School, Federal University of Rio de Janeiro, RJ, Brazil.
Running head: Anticariogenic effect of a Brazilian green propolis Correspondence to: Andréa Gonçalves Antonio Rua Professor Coutinho Fróis 500/301, Rio de Janeiro – RJ, Brazil, Zip Code: 22620360, e-mail: [email protected]
2
Highlights
The inhibitory action of a Wild green propolis was tested against a S. mutans dental biofim.
The tested propolis inhibited the growth of S. mutans biofilm.
The tested propolis showed low level of influence in inhibiting the demineralization of the enamel analyzed.
Abstract Objective: This study investigated the anti-demineralizing and antibacterial effects of a propolis ethanolic extract (EEP) against Streptococcus mutans dental biofilm. Design: Blocks of sound bovine enamel (n=24) were fixed on polystyrene plates. S. mutans inoculum (ATCC 25175) and culture media were added (48h–37ºC) to form biofilm. Blocks with biofilm received daily treatment (30μL/1min), for 5 days, as following: G1 (EEP 33.3%); G2 (chlorhexidine digluconate 0.12%); G3 (ethanol 80%); and G4 (MilliQ water). G5 and G6 were blocks without biofilm that received only EEP and Milli-Q water, respectively. Final surface hardness was evaluated and the percentage of hardness loss (%HL) was calculated. The EEP extract pH and total solids were determined. S. mutans count was expressed by log10 scale of Colony-Forming Units (CFU/mL). One way ANOVA was used to compare results which differed at a 95% significance level.
Results:
G2
presented the
lowest
average
%HL
value
(68.44%±12.98) (p=0.010), while G4 presented the highest (90.49%±5.38 %HL) (p=0.007). G1 showed %HL (84.41%±2.77) similar to G3 (87.80%±6.89) (p=0.477). Groups G5 and G6 presented %HL=16.11%±7.92 and 20.55%±10.65; respectively (p=0.952). G1 and G4 differed as regards to S. mutans count: 7.26±0.08 and 8.29±0.17 CFU/mL, respectively (p=0.001). The lowest bacterial count was observed in chlorhexidine group (G2=6.79±0.10 CFU/mL) (p=0.043).There was no difference between S. mutans count of G3 and G4 (p=0.435). The EEP showed pH=4.8 and total soluble solids content=25.9 °Brix.
Conclusion: The EEP seems to be a potent
antibacterial substance against S. mutans dental biofilm, but presented no inhibitory action on the de-remineralization of caries process. Keywords: propolis; antibacterial activity; dental biofilm; Streptococcus mutans; microhardness
3 Introduction Dental caries is a condition that depends on the presence of biofilm or dental plaque Cury & Tenuta, 2009). When the microbial deposits remain adhered to the tooth, episodes of pH drop in the biofilm exposed to sugar occur (Filoche, Wong, & Sissons, 2010) leading to a loss of enamel integrity (Takahashi & Nyvad, 2008). Therefore, in face of a diet rich in sugar and fermentable carbohydrates, the pH of plaque remains low, favoring the cariogenic microbiota growth, which consequently increases the risk for caries disease (Kidd & Fejerskov, 2004). In this sense, some studies (Touger-Decker & Van Loveren, 2003; Bonow, Azevedo, Goettems, & Rodrigues, 2013) have been conducted with the purpose to determine anticariogenic strategies to reduce the risks posed by sugar and other fermentable carbohydrates, exploring the use of natural products. These products may prevent caries disease without provoke undesirable side effects such as development of bacteria to tolerance, vomiting, diarrhea and others (Chandra Shekar, Nagarajappa, Suma, & Thakur, 2015). Among these products, different types of plants rich in polyphenols, such as coffee, green tea, cocoa and grape, have been investigated with regard to their antibacterial action against caries pathogens (Ooshima et al., 2000; Smullen et al., 2007; Antonio et al., 2011; Yano et al., 2012; Meckelburg et al., 2014). Propolis, also rich in polyphenols (Koo et al., 2002), has aroused scientific interest with regard to its beneficial pharmacological properties to oral health, such as the anti-inflammatory and antimicrobic (Eick et al., 2014; Abubakar, Abdullah, Sulaiman, & Ang, 2014). Additionally, the effective concentrations of mouthrinse containing propolis on oral microorganisms were less cytotoxic on human gingival fibroblasts than Chlorhexidine (Ozan et al., 2007). However, there are still gaps to be elucidated considering its effect on the de-remineralization process of caries. Bearing in mind that the development of new therapies with the use of natural products for prevention and treatment of diseases is relevant in the field of medicine (Walker, 1996), this study aims to investigate the anti-demineralizing and antibacterial effects of a propolis extract against Streptococcus mutans biofilm formed on bovine dental enamel.
4 Materials and Methods Propolis Ethanolic extract Sample: Wild green propolis was collected from Apis mellifera beehives in the Atlantic Rain Forest of Bocaina mountains, town of São José do Barreiro, State of São Paulo, Brazil (latitude 22°45’ South, longitude 44° 39’ West of Greenwich, 1200m altitude) and kept in a desiccator for a week. Crude propolis was ground in nitrogen and sieved through a 0.6 mm sieve. The extract was prepared according to Park et al. (1998), modified as follows: 30mL of 80% ethanol were added to 10g of ground propolis. The mixture was heated to 60 °C for 30 min., under agitation, filtered through Watman n.2 paper and centrifuged at 7500 x g at 5 °C for 10 min. The supernatant (propolis extract – EEP) was directly used for the biofilm assay. pH, total solids The pH was determined with a pH meter (DM20 Digitized, Santo Amparo, SP, Brazil). Total soluble solids were determined using a digital refractometer (ATAGO®, PAL-1, Japan) and results were expressed in °Brix. Determination of total Polyphenol content Total polyphenols were measured by colorimetric assays using the FolinCiocalteu method (Singleton, Orthofer, & Lamuela-Raventos, 1999) with modifications. Each extract was diluted 1:10 in distilled water; then 30 µL of this dilution was mixed with 120 µL of distilled water, 75 µL of Folin-Ciocalteu reagent (Merck, Germany) and 75 µL sodium carbonate 20% (w/v). The absorbance was measured at 760 nm (UV1800) after 30 min incubation at room temperature. The concentrations were calculated using a calibration curve and were expressed in mg/mL gallic acid equivalent Determination of Hydroxycinnamic acids and derivatives The contents of caffeic, ferulic, p-coumaric and 5-caffeoylquinic acids (from SIGMA-Aldrich, Germany) and 3-caffeoylquinic, 4-caffeoylquinic,3,4-dicafeoylquinic, 3,5-dicaffeoylquinic, 4,5-dicaffeoylquinic (IUPAC numbering), were investigated by HPLC-DAD-reverse-phase system, with clarification and chromatographic conditions based on Farah et al. (2005). Bacterial Strain and Inoculum Preparation A bacterial sample of Streptococcus mutans (ATCC 25175) was used to prepare the inoculum. Initially, the bacterial sample was evaluated to verify the degree of purity. After this, isolated bacterial colonies were selected and transferred to a 0.85% saline solution until an Optical Density (O.D.) of 0.15 at 520nm (Libra S2 Colorimeter, Biochrom, Cambridge, England) corresponding to approximately 108 CFU/mL, was obtained. Determination of Minimum Inhibitory Concentration (MIC) and Minimum
5 Bactericidal Concentration (MBC) The antibacterial activity of EEP was examined inicially, by determining the minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC), according to the Clinical and Laboratory Standards Institute (CLSI) guidelines (CLSI, 2012) with a modification proposed by Cunha et al. (2013). MIC was performed in 96-well microplates, inoculated with 5 X 105 CFU/mL, in 100 µL of brain heart infusion medium (BHI, Oxoid Ltd., Hampshire, England). The concentrations of EEP ranged from 266.67 to 0.259 mg/mL. Plain ethanol solution was used as the vehicle control (concentrations ranged from 64 to 0.0625%, (v/v)). Chlorhexidine digluconate 0.05% was used as positive control. Plates were incubated at 37°C and 5% CO2 for 24 h and MIC was defined as the lowest concentration of EEP that allowed no visible growth, confirmed by 0.01% resazurin dye (Sigma-Aldrich,St. Louis, MO, USA). The wells with the highest concentration of EEP and visible growth after the incubation were analyzed for purity of the bacterial suspension by surface culture on BHI agar (Oxoid Ltd., Hampshire, England). MBC was determined by subculturing 50µL aliquots of each incubated well that presented concentration higher than the MIC on BHI agar. Two separate experiments were conducted in triplicate for each concentration of EEP. Selection and preparation of bovine tooth samples Fifty four sound bovine incisors, disinfected in 2% formol (pH 7.0), were selected after confirming absence of caries, stains, cracks or other enamel defects. They were cut using a water-cooled diamond saw (Bühler, Uzwil, Switzerland) to obtain enamel blocks (4 x 4 x 2 mm). These blocks were fixed with wax in an acrylic device to polish the enamel surface: 600- and 1200-grit silicon carbide papers (Extec Corp., Enfield, USA), followed by 1-µm diamond abrasive slurry (Extec Corp., Enfield, USA) and washed ultrasonically in Milli-Q water (Merck Millipore, Darmstadt, Germany). Following, the specimens were stored in a physiological solution for posterior analysis of surface hardness. Initial surface hardness evaluation and selection of enamel blocks for test with biofilm
The surface microhardness test was performed using a Microhardness Tester (Micromet 5104; Buehler, Mitutoyo Corporation, Tokyo, Japan) with a Knoop type diamond under a 50g load for 5 seconds. Three indentations spaced 100 µm from each other were made at the center of the enamel surface to select the sample (Mazer Papa et al., 2010). The mean of these three values represented the sample hardness.
6 Twenty four blocks (mean 330.00 ± 31.15 kg/mm2) were used in the present study. All samples were stored in a humid environment. In vitro S. mutans biofilm formation on bovine tooth fragments The experimental protocol was carried out in accordance with the methodology proposed by Soares et al. (2015). Each bovine tooth fragment (n = 24) was fixed with wax in a single well in a 24-well polystyrene plate. This plate/fragment system was sterilized with ethylene oxide (Bioxxi, Serviços de Esterilização LTDA, Rio de Janeiro, Brazil). After sterilization, culture medium (BHI supplemented with 2% sucrose, Oxoid Ltd, 1492.5 µL/well) was added to each well together with the bacterial inoculum (7.5 µL of S. mutans), thus representing an inoculum with a final concentration of approximately 5 x 105 CFU/mL. Once this procedure was carried out, the system was incubated under microaerofilia at 37ºC for 48 hours, so that a mature biofilm would form on the tooth fragments. Treatment of biofilm After biofilm formation, the fragments were divided according to the following treatment groups: G1 (EEP 33.33%; n=4); G2 (0.12% Chlorhexidine, n=4); G3 (80% Ethanol; n=4); G4 (Milli-Q Water; n=4). Eight blocks without biofilm were treated with EEP (G5, n=4) and Milli-Q water (G6, n=4). Treatment was performed according to the following protocol: the culture medium was removed from each well, and after this, the surface of each specimen with (G1, G2, G3, G4) and without (G5, G6) biofilm received 30 μL of the test substance, which remained on the fragments for one minute. The specimens were then washed three times with 1000 μL of sterile Milli-Q water, and a new culture medium (BHI, Difco, 1500 μL), without the inoculum, was added to the well. This procedure was repeated every 24 hours. The plate/specimen/biofilm system was again incubated under microaerofilia, in which a new application of the test substance was performed (always at the same time), for the period of 5 consecutive days. Evaluation of final surface microhardness of bovine enamel After microbiological analyses of the biofilm, the fragments were vortexed until they were completely cleaned, and prepared once more for the final surface microhardness test. The same surface microhardness test parameters were followed. Three new indentations were made, 150µm equidistant from the first markings (initial indentations) and also spaced at 100µm from one another (Cunha et al., 2013).The mean of these three values represented the final hardness of the sample. The percentage hardness loss was calculated in compliance with the following expression: (initial hardness – final hardness/ initial hardness) × 100.
7 This test was performed by a blind examiner. Microbiological analysis of the S. mutans biofilm formed on the bovine tooth fragments After five days of the experiment, the bovine enamel fragments were inserted into tubes containing 1mL of saline solution. Following, the tube/fragment system was vortexed for 2 minutes, in order to detach the biofilm from the tooth. From this suspension, aliquots of 50 µL were removed for seeding (dilutions of 10 -1 to 10-7), in duplicate, on petri dishes containing BHI agar (Difco) for S. mutans counts. Plates were incubated at 37ºC and 5% CO2, for 48h and the results were expressed by log10 scale of Colony-Forming Units (CFU/mL). Statistical Analyses Data were analyzed by means of the statistical software program SPSS version 20.0 (SPSS Inc., Chicago, USA). The Shapiro-Wilk test was used to verify the distribution of normality of the microbiologic results and of the percentage of hardness loss of the bovine tooth enamel blocks. With regard to the microbial count and the hardness loss values, ANOVA followed by Tukey test was used to verify whether there was statistical difference among the groups of treatment. Sample power was calculated between the groups EEP (tested substance) and the Chlorhexidine (positive control) using an α error of 5%, which resulted in a power of 0.8752.
Results The Hydroxycinnamic acids and derivatives content of the EEP at 33.33%, as well as the total phenolic compound content are expressed in Table 1. The pH of the EEP was 4.8 and the total solid content was 25.9 °Brix. The minimum inhibitory concentration of EEP against S. mutans was 2.08 mg/mL, and 8% for ethanol; while the minimum bactericide concentration of the extract and ethanol was 8.33mg/mL and 16%, respectively. After treating the biofilm formed on the bovine enamel fragments, G1 (EEP) percentage of hardness loss (%HL) was 84.41 ± 2.77, similar (p = 0.477) to G3 (Ethanol) (%HL = 87.80 ± 6.89). The %HL value (68.44 ± 12.98) of G2 (Chlorhexidine) was the lowest in comparison with the other groups (p = 0.010). G4 (Milli-Q water) presented the highest mean value for %HL (90.49±5.38) after the period of treatment. The groups G5 and G6, treated with EEP and Milli-Q water, respectively, but with no microorganisms, presented the following %HL: 16.11% ± 7.92; and 20.55% ± 10.65; respectively (p=0.952). All the results with reference to hardness loss of fragments may be visualized in Table 2. There was difference between G1 (EEP) and G4 (Milli-Q water) as regards the
8 number of S. mutans: 7.26±0.08 and 8.29±0.17CFU/mL, respectively (p = 0.001). The lower quantity of viable cells was presented by G2 (Chlorhexidine) (6.79±0.10 CFU/mL) (p = 0.043), and there was no difference (p = 0.435) between G3 and G4 (Table 2).
Discussion There are many strategies that can be implemented to control dental caries, such as: a low sugar diet, instruction to oral hygiene, use of fluoride dentifrice, the control of biofilm, and others. Besides that, there are evidences (Chandra Shekar, Nagarajappa, Suma, & Thakur, 2015) that support the advantages of natural products as coadjutant factors to be used in specific cases of caries activity. According to Chandra-Shekar, Nagarajappa, Suma, & Thakur (2015), the use of herbal extracts for the control of oral/dental diseases is considered to be an interesting alternative to synthetic antimicrobials due to their lower negative impact and to overcome intrinsic (primary) resistance or secondary resistance to drugs during therapy. Therefore, in an effort to reduce the prevalence of caries, different natural products have been previously tested for anticariogenic effect (Cheng, Li, He, & Zhou, 2015; Almeida et al., 2012). It is important to emphasize that the anticariogenic effect of a product involves both inhibition of growth of the bacteria responsible for caries disease, and inhibition of the enamel demineralization process (Antonio et al., 2011); effects tested in the present study. It is known that there is not only one bacterial species involved in the carious process (Kidd & Fejerskov, 2004; Gross et al., 2012). However, although Streptococcus mutans is not the only microorganism responsible for the onset of the disease, the biofilm formed by these species is usually chosen to evaluate the antibacterial and/or anticariogenic activities of different products (Bueno-Silva et al., 2013; Fernández, Giacaman, Tenuta, & Cury, 2015). According to Bueno-Silva et al. (2013), Streptococcus mutans and sucrose are key modulators of the development of cariogenic biofilms. Considering the antibacterial effect of phenolic compounds, studies have pointed out that these substances are the main active ingredients of natural products responsible for this function. They inhibit the enzyme glycosyltransferase, responsible among other factors, for the adherence of S. mutans to the tooth surface for dental biofilm formation (Koo et al., 2002). In the present study, the total polyphenol and hydroxycinnamic acids contents found in propolis was low (Table 1) in comparison with data from previous studies (Souza et al., 2007; Cabral et al., 2009). In spite of this finding, it is probable that the presence of caffeic and 5-caffeoylquinic acids (5-CQA) found in the tested extract, in synergy with other possible compounds, may have
9 contributed to decrease the number of viable microorganisms observed after treatment of the blocks/biofilm with EEP. According to Antonio et al. (2010), 5-CQA and caffeic acid are substances capable of inhibiting the growth of S. mutans. The microorganisms count in the group treated with EEP was significantly lower than that of the groups treated with 8% ethanol and Milli-Q water (Table 2). This result shows that EEP was efficient in reducing the number of S. mutans present in dental biofilm. However, the mean of the hardness loss percentage in the group (G1) treated with EEP (%HL = 84.41% ± 2.77) did not differ statistically from that of the groups treated with Milli-Q water (G4) and Ethanol (G3). According to Zhang et al. (2009), the enamel demineralization process may be inhibited by the interaction of polyphenols with the organic matrix, which results in lower mineral loss. These interactions involve covalent, ionic, and hydrogen bonding or hydrophobic processes, which induce the metamorphism of the enamel organic matrix. The metamorphic organic matrix is precipitated on the enamel, resulting in slowing down the speed of mineral ion loss, and consequently, enamel demineralization is inhibited. We did not find some of the phenolic components investigated (Table 1) and, consequently, a low variety of polyphenols of the propolis tested was observed. It possibly explains the low level of influence of this product in inhibiting the demineralization of the enamel analyzed, in spite of its capacity to act directly on the bacteria in conjunction with the other compounds, but the presence of other types of active polyphenols such as flavonoids, which were not individually analyzed in this study, is possible. This controversial result leads to the believe that the time of treatment may have played a fundamental role in this case, because lysis of the bacteria may have occurred in a period after the loss of hardness already in progress in the enamel; and the acidic pH (4.8) of the propolis extract may also have contributed to these results. Furthermore, no tests were performed with mixed biofilm and the authors suggest further studies on this subject. Also, although the power of the study presented a favorable result between the EEP and Chlorhexidine groups, the authors believe that a large sample size is needed in future studies. Another point to consider is the high concentration (80%) of ethanol that was used for biofilm treatment, which may have contributed to a smaller number of bacteria acting on dental demineralization, and therefore, to the results of hardness loss in G3 which, although higher, did not differ from those of G1. The minimum inhibitory concentration (MIC) of the substance was 2.08 mg/mL and the bactericidal concentration (MBC), 8.33 mg/mL. The values found are higher than those of previous studies (Cabral et al., 2009; Castro et al., 2009; Swerts, Costa, & Fiorini, 2010). The chemical composition of propolis is variable, according to its type,
10 origin and seasonality, which leads to its diverse biologic effects, such as antiinflammatory, antimicrobial and antioxidant, as shown in previous studies (Akao et al., 2003; Cabral et al., 2009). Therefore, this fact perfectly justifies the continuous study of propolis. For a better qualitative study of the extract as regards to its consumption, cytotoxicity tests must be conducted, which represents a limitation of the present study. However, in accordance with the administrative ruling of the Ministry of Agriculture of Brazil (Brasil, 2015), the technical standard for determination of identity and quality of propolis for consumption reads that the minimum content of phenolic compounds must be 0.50%. Thus, the contents found in the extract analyzed in the present study were above this value. Moreover, the cariogenic challenge with sucrose concentrations was constant up to the end of the experimental protocol, which also represents a limitation of this study. However, according to Pierro et al. (2014) a sudden exposure to an excess amount of sugar in laboratory studies may cause bacterial death because the rapid entry and degradation of sugars in cells can result in an accumulation of toxic levels of glycolytic intermediates. Thus, taking into consideration that it was an in vitro study, the authors only opted to frequently change the media to maintain the viability of the microorganisms. A previous study (Duarte et al., 2006) investigated the ethanolic extract of a novel type of propolis and its purified hexane fraction on S. mutans biofilms and the development of dental caries in rats. The results suggested that the cariostatic properties of propolis were due to the reduction of acid production and acid tolerance of cariogenic streptococci (Duarte et al., 2006). Recent clinical studies (Mohsin, Manohar, Rajesh, & Asif, 2015; Prabhakar, Karuna, Yavagal, & Deepak, 2015) reinforced the role of propolis, in their extract form or incorporated in dentifrices, to reducethe microbial load throughout the human oral cavity. Also, with the results obtained in the present research, the authors suggest that this propolis extract is a potent inhibitor of S. mutans biofilm but it was not able to inhibit the demineralization process of the dental enamel. However, in the future, comparison of the anticariogenic activity of different types of propolis, may possibly predict the most indicated types for use in dentistry.
Acknowledgments The authors thank the Fundação de Amparo à Pesquisa do Estado do Rio de Janeiro (FAPERJ) for the financial support and Sérgio Lutz Barbosa for the green própolis sample.
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Pierro VS, Iorio NL, Lobo LA, Cabral LM, Dos Santos KR, Maia LC. Effect of a sugar-free pediatric antibiotic on primary tooth enamel hardness when exposed to different sucrose exposure conditions in situ. Clin Oral Investig 2014; 18(5):1391-9.
Prabhakar AR, Karuna YM, Yavagal C, Deepak BM. Cavity disinfection in minimally invasive dentistry - comparative evaluation of Aloe vera and propolis: A randomized clinical trial. Contemp Clin Dent 2015; 6 (Suppl 1):S24-31.
Santos FA, Bastos EM, Uzeda M, Carvalho MA, Farias LM, Moreira ES, Braga FC. Antibacterial activity of Brazilian propolis and fractions against oral anaerobic bacteria. J Ethnopharmacol 2002; 80(1):1-7.
14
Singleton VL, Orthofer R, Lamuela-Raventos RM. Analysis of total phenols and other oxidation substrates and antioxidants by means of Folin-Ciocalteu reagent. Methods Enzymol 1999; 299: 152-78.
Smullen J, Koutsou GA, Foster HA, Zumbé A Storey DM. The antibacterial activity of plant extracts containing polyphenols against Streptococcus mutans. Caries Res 2007; 41(5):342-49.
Soares DN, Antonio AG, Iorio NL, Pierro VS, dos Santos KR, Maia LC. Does the presence of sucrose in pediatric antibiotics influence the enamel mineral loss and the Streptococcus mutans counts in dental biofilm? Braz Dent J 2015; 26(3):249-57.
Sousa JP, Furtado NAJC, Jorge R, Soares AE, Bastos JK. Perfis físico-químico e cromatográfico de amostras de própolis produzidas nas microrregiões de Franca (SP) e Passos (MG), Brasil. Rev Bras Farmacogn 2007; 17(1):85-93.
Swerts MSO, Costa AMDD, Fiorini JE. Efeito da Solução Associada de Clorexidina e Própolis na Inibição da Aderência de Streptococcus spp. Rev Int Periondontia Clínica 2010; 2(4):10-6.
Takahashi N, Nyvad B. Caries ecology revisited: microbial dynamics and the caries process. Caries Res 2008; 42(6):409-18.
Touger-Decker R, Van Loveren, C. Sugars and dental caries. Am J Clin Nutr 2003; 78(4):881S-892S.
Walker CB. The acquisition of antibiotic resistance in the periodontal microflora. Periodontol 2000 1996; 10:79-88.
Yano A, Kikuchi S, Takahashi T, Kohama K, Yoshida Y. Inhibitory effects of the phenolic fraction from the pomace of Vitis coignetiae on biofilm formation by Streptococcus mutans. Arch Oral Biol 2012; 57(6):711-9.
Zhang L, Xue J, Li J, Zou L, Hao Y, Zhou X, Li W. Effects of Galla chinensis on inhibition of demineralization of regular bovine enamel or enamel disposed of organic matrix. Arch Oral Biol 2009; 54(9):817-22.
15 Table 1: Hydroxycinnamic acids and derivatives presented in the ethanol extract of propolis Hydroxycinnamic acids and derivatives
(µg/mL)
3-caffeoylquinic acid
0.0
4-caffeoylquinic acid
4.84
5-caffeoylquinic acid
14.6
Caffeic acid
23.5
Ferulic acid
0.0
p-coumaric acid
0.0
3,4-dicaffeoylquinic acid
27.3
3,5-dicaffeoylquinic acid
18.4
4,5-dicaffeoylquinic acid
0.0
Total
88.64
16 Table 2: Percentage hardness loss (%) and number of Streptococcus mutans colonies present in biofilm after treatment of different studied groups (G) S. mutans (CFU/biofilm)
Group
Hardness loss (%)
G1
84.41±2.77a
7.26±0.08
G2
68.44±12.98b
6.79±0.10
G3
87.80±6.89a
8.00±0.05
G4
90.49±5.38a
8.29±0.17
a
b
c
c
Note: Study Groups: G1 – 33.33% EEP; G2 – 0.12% Chlorhexidine; G3 – 80% Ethanol c/v EEP; G4 – Milli-Q water. Different letters in the same column represent statistically significant results. Results of the microbial count are shown in log10 scale for better comparison.
S0003-9969(16)30028-0 http://dx.doi.org/doi:10.1016/j.archoralbio.2016.02.001 AOB 3544
To appear in:
Archives of Oral Biology
Received date: Revised date: Accepted date:
29-5-2015 19-12-2015 1-2-2016
Please cite this article as: Cardoso Julia Gabiroboertz, Iorio Natalia Lopes Pontes, Rodrigues Lu´is Fernando, Couri Maria Luiza Barra, Farah Adriana, Maia Lucianne Cople, Antonio Andr´ea Gonc¸alves.Influence of a Brazilian wild green propolis on the enamel mineral loss and Streptococcus mutans’ count in dental biofilm.Archives of Oral Biology http://dx.doi.org/10.1016/j.archoralbio.2016.02.001 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
1 Influence of a Brazilian wild green propolis on the enamel mineral loss and Streptococcus mutans’ count in dental biofilm Julia Gabiroboertz Cardoso1, Natalia Lopes Pontes Iorio2, Luís Fernando Rodrigues3, Maria Luiza Barra Couri4, Adriana Farah5, Lucianne Cople Maia6, Andréa Gonçalves Antonio7 1 Graduate
student, Department of Pediatric Dentistry and Orthodontics, Dental School,
Federal University of Rio de Janeiro, RJ, Brazil. 2
PhD, Adjunct Professor, Department of Basic Science, Federal Fluminense
University, Nova Friburgo Campus, Brazil. 3
Graduate student, Department of Basic Science, Federal Fluminense University,
Nova Friburgo Campus, Brazil. 4
Graduate student, Department of Basic Science, Federal Fluminense University,
Nova Friburgo Campus, Brazil. 5 PhD,
Adjunct Professor, Institute of Nutrition, Federal University of Rio de Janeiro, RJ,
Brazil. 6
PhD, Full Professor, Department of Pediatric Dentistry and Orthodontics, Dental
School, Federal University of Rio de Janeiro, RJ, Brazil. 7
PhD, Adjunct Professor, Department of Pediatric Dentistry and Orthodontics, Dental
School, Federal University of Rio de Janeiro, RJ, Brazil.
Running head: Anticariogenic effect of a Brazilian green propolis Correspondence to: Andréa Gonçalves Antonio Rua Professor Coutinho Fróis 500/301, Rio de Janeiro – RJ, Brazil, Zip Code: 22620360, e-mail: [email protected]
2
Highlights
The inhibitory action of a Wild green propolis was tested against a S. mutans dental biofim.
The tested propolis inhibited the growth of S. mutans biofilm.
The tested propolis showed low level of influence in inhibiting the demineralization of the enamel analyzed.
Abstract Objective: This study investigated the anti-demineralizing and antibacterial effects of a propolis ethanolic extract (EEP) against Streptococcus mutans dental biofilm. Design: Blocks of sound bovine enamel (n=24) were fixed on polystyrene plates. S. mutans inoculum (ATCC 25175) and culture media were added (48h–37ºC) to form biofilm. Blocks with biofilm received daily treatment (30μL/1min), for 5 days, as following: G1 (EEP 33.3%); G2 (chlorhexidine digluconate 0.12%); G3 (ethanol 80%); and G4 (MilliQ water). G5 and G6 were blocks without biofilm that received only EEP and Milli-Q water, respectively. Final surface hardness was evaluated and the percentage of hardness loss (%HL) was calculated. The EEP extract pH and total solids were determined. S. mutans count was expressed by log10 scale of Colony-Forming Units (CFU/mL). One way ANOVA was used to compare results which differed at a 95% significance level.
Results:
G2
presented the
lowest
average
%HL
value
(68.44%±12.98) (p=0.010), while G4 presented the highest (90.49%±5.38 %HL) (p=0.007). G1 showed %HL (84.41%±2.77) similar to G3 (87.80%±6.89) (p=0.477). Groups G5 and G6 presented %HL=16.11%±7.92 and 20.55%±10.65; respectively (p=0.952). G1 and G4 differed as regards to S. mutans count: 7.26±0.08 and 8.29±0.17 CFU/mL, respectively (p=0.001). The lowest bacterial count was observed in chlorhexidine group (G2=6.79±0.10 CFU/mL) (p=0.043).There was no difference between S. mutans count of G3 and G4 (p=0.435). The EEP showed pH=4.8 and total soluble solids content=25.9 °Brix.
Conclusion: The EEP seems to be a potent
antibacterial substance against S. mutans dental biofilm, but presented no inhibitory action on the de-remineralization of caries process. Keywords: propolis; antibacterial activity; dental biofilm; Streptococcus mutans; microhardness
3 Introduction Dental caries is a condition that depends on the presence of biofilm or dental plaque Cury & Tenuta, 2009). When the microbial deposits remain adhered to the tooth, episodes of pH drop in the biofilm exposed to sugar occur (Filoche, Wong, & Sissons, 2010) leading to a loss of enamel integrity (Takahashi & Nyvad, 2008). Therefore, in face of a diet rich in sugar and fermentable carbohydrates, the pH of plaque remains low, favoring the cariogenic microbiota growth, which consequently increases the risk for caries disease (Kidd & Fejerskov, 2004). In this sense, some studies (Touger-Decker & Van Loveren, 2003; Bonow, Azevedo, Goettems, & Rodrigues, 2013) have been conducted with the purpose to determine anticariogenic strategies to reduce the risks posed by sugar and other fermentable carbohydrates, exploring the use of natural products. These products may prevent caries disease without provoke undesirable side effects such as development of bacteria to tolerance, vomiting, diarrhea and others (Chandra Shekar, Nagarajappa, Suma, & Thakur, 2015). Among these products, different types of plants rich in polyphenols, such as coffee, green tea, cocoa and grape, have been investigated with regard to their antibacterial action against caries pathogens (Ooshima et al., 2000; Smullen et al., 2007; Antonio et al., 2011; Yano et al., 2012; Meckelburg et al., 2014). Propolis, also rich in polyphenols (Koo et al., 2002), has aroused scientific interest with regard to its beneficial pharmacological properties to oral health, such as the anti-inflammatory and antimicrobic (Eick et al., 2014; Abubakar, Abdullah, Sulaiman, & Ang, 2014). Additionally, the effective concentrations of mouthrinse containing propolis on oral microorganisms were less cytotoxic on human gingival fibroblasts than Chlorhexidine (Ozan et al., 2007). However, there are still gaps to be elucidated considering its effect on the de-remineralization process of caries. Bearing in mind that the development of new therapies with the use of natural products for prevention and treatment of diseases is relevant in the field of medicine (Walker, 1996), this study aims to investigate the anti-demineralizing and antibacterial effects of a propolis extract against Streptococcus mutans biofilm formed on bovine dental enamel.
4 Materials and Methods Propolis Ethanolic extract Sample: Wild green propolis was collected from Apis mellifera beehives in the Atlantic Rain Forest of Bocaina mountains, town of São José do Barreiro, State of São Paulo, Brazil (latitude 22°45’ South, longitude 44° 39’ West of Greenwich, 1200m altitude) and kept in a desiccator for a week. Crude propolis was ground in nitrogen and sieved through a 0.6 mm sieve. The extract was prepared according to Park et al. (1998), modified as follows: 30mL of 80% ethanol were added to 10g of ground propolis. The mixture was heated to 60 °C for 30 min., under agitation, filtered through Watman n.2 paper and centrifuged at 7500 x g at 5 °C for 10 min. The supernatant (propolis extract – EEP) was directly used for the biofilm assay. pH, total solids The pH was determined with a pH meter (DM20 Digitized, Santo Amparo, SP, Brazil). Total soluble solids were determined using a digital refractometer (ATAGO®, PAL-1, Japan) and results were expressed in °Brix. Determination of total Polyphenol content Total polyphenols were measured by colorimetric assays using the FolinCiocalteu method (Singleton, Orthofer, & Lamuela-Raventos, 1999) with modifications. Each extract was diluted 1:10 in distilled water; then 30 µL of this dilution was mixed with 120 µL of distilled water, 75 µL of Folin-Ciocalteu reagent (Merck, Germany) and 75 µL sodium carbonate 20% (w/v). The absorbance was measured at 760 nm (UV1800) after 30 min incubation at room temperature. The concentrations were calculated using a calibration curve and were expressed in mg/mL gallic acid equivalent Determination of Hydroxycinnamic acids and derivatives The contents of caffeic, ferulic, p-coumaric and 5-caffeoylquinic acids (from SIGMA-Aldrich, Germany) and 3-caffeoylquinic, 4-caffeoylquinic,3,4-dicafeoylquinic, 3,5-dicaffeoylquinic, 4,5-dicaffeoylquinic (IUPAC numbering), were investigated by HPLC-DAD-reverse-phase system, with clarification and chromatographic conditions based on Farah et al. (2005). Bacterial Strain and Inoculum Preparation A bacterial sample of Streptococcus mutans (ATCC 25175) was used to prepare the inoculum. Initially, the bacterial sample was evaluated to verify the degree of purity. After this, isolated bacterial colonies were selected and transferred to a 0.85% saline solution until an Optical Density (O.D.) of 0.15 at 520nm (Libra S2 Colorimeter, Biochrom, Cambridge, England) corresponding to approximately 108 CFU/mL, was obtained. Determination of Minimum Inhibitory Concentration (MIC) and Minimum
5 Bactericidal Concentration (MBC) The antibacterial activity of EEP was examined inicially, by determining the minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC), according to the Clinical and Laboratory Standards Institute (CLSI) guidelines (CLSI, 2012) with a modification proposed by Cunha et al. (2013). MIC was performed in 96-well microplates, inoculated with 5 X 105 CFU/mL, in 100 µL of brain heart infusion medium (BHI, Oxoid Ltd., Hampshire, England). The concentrations of EEP ranged from 266.67 to 0.259 mg/mL. Plain ethanol solution was used as the vehicle control (concentrations ranged from 64 to 0.0625%, (v/v)). Chlorhexidine digluconate 0.05% was used as positive control. Plates were incubated at 37°C and 5% CO2 for 24 h and MIC was defined as the lowest concentration of EEP that allowed no visible growth, confirmed by 0.01% resazurin dye (Sigma-Aldrich,St. Louis, MO, USA). The wells with the highest concentration of EEP and visible growth after the incubation were analyzed for purity of the bacterial suspension by surface culture on BHI agar (Oxoid Ltd., Hampshire, England). MBC was determined by subculturing 50µL aliquots of each incubated well that presented concentration higher than the MIC on BHI agar. Two separate experiments were conducted in triplicate for each concentration of EEP. Selection and preparation of bovine tooth samples Fifty four sound bovine incisors, disinfected in 2% formol (pH 7.0), were selected after confirming absence of caries, stains, cracks or other enamel defects. They were cut using a water-cooled diamond saw (Bühler, Uzwil, Switzerland) to obtain enamel blocks (4 x 4 x 2 mm). These blocks were fixed with wax in an acrylic device to polish the enamel surface: 600- and 1200-grit silicon carbide papers (Extec Corp., Enfield, USA), followed by 1-µm diamond abrasive slurry (Extec Corp., Enfield, USA) and washed ultrasonically in Milli-Q water (Merck Millipore, Darmstadt, Germany). Following, the specimens were stored in a physiological solution for posterior analysis of surface hardness. Initial surface hardness evaluation and selection of enamel blocks for test with biofilm
The surface microhardness test was performed using a Microhardness Tester (Micromet 5104; Buehler, Mitutoyo Corporation, Tokyo, Japan) with a Knoop type diamond under a 50g load for 5 seconds. Three indentations spaced 100 µm from each other were made at the center of the enamel surface to select the sample (Mazer Papa et al., 2010). The mean of these three values represented the sample hardness.
6 Twenty four blocks (mean 330.00 ± 31.15 kg/mm2) were used in the present study. All samples were stored in a humid environment. In vitro S. mutans biofilm formation on bovine tooth fragments The experimental protocol was carried out in accordance with the methodology proposed by Soares et al. (2015). Each bovine tooth fragment (n = 24) was fixed with wax in a single well in a 24-well polystyrene plate. This plate/fragment system was sterilized with ethylene oxide (Bioxxi, Serviços de Esterilização LTDA, Rio de Janeiro, Brazil). After sterilization, culture medium (BHI supplemented with 2% sucrose, Oxoid Ltd, 1492.5 µL/well) was added to each well together with the bacterial inoculum (7.5 µL of S. mutans), thus representing an inoculum with a final concentration of approximately 5 x 105 CFU/mL. Once this procedure was carried out, the system was incubated under microaerofilia at 37ºC for 48 hours, so that a mature biofilm would form on the tooth fragments. Treatment of biofilm After biofilm formation, the fragments were divided according to the following treatment groups: G1 (EEP 33.33%; n=4); G2 (0.12% Chlorhexidine, n=4); G3 (80% Ethanol; n=4); G4 (Milli-Q Water; n=4). Eight blocks without biofilm were treated with EEP (G5, n=4) and Milli-Q water (G6, n=4). Treatment was performed according to the following protocol: the culture medium was removed from each well, and after this, the surface of each specimen with (G1, G2, G3, G4) and without (G5, G6) biofilm received 30 μL of the test substance, which remained on the fragments for one minute. The specimens were then washed three times with 1000 μL of sterile Milli-Q water, and a new culture medium (BHI, Difco, 1500 μL), without the inoculum, was added to the well. This procedure was repeated every 24 hours. The plate/specimen/biofilm system was again incubated under microaerofilia, in which a new application of the test substance was performed (always at the same time), for the period of 5 consecutive days. Evaluation of final surface microhardness of bovine enamel After microbiological analyses of the biofilm, the fragments were vortexed until they were completely cleaned, and prepared once more for the final surface microhardness test. The same surface microhardness test parameters were followed. Three new indentations were made, 150µm equidistant from the first markings (initial indentations) and also spaced at 100µm from one another (Cunha et al., 2013).The mean of these three values represented the final hardness of the sample. The percentage hardness loss was calculated in compliance with the following expression: (initial hardness – final hardness/ initial hardness) × 100.
7 This test was performed by a blind examiner. Microbiological analysis of the S. mutans biofilm formed on the bovine tooth fragments After five days of the experiment, the bovine enamel fragments were inserted into tubes containing 1mL of saline solution. Following, the tube/fragment system was vortexed for 2 minutes, in order to detach the biofilm from the tooth. From this suspension, aliquots of 50 µL were removed for seeding (dilutions of 10 -1 to 10-7), in duplicate, on petri dishes containing BHI agar (Difco) for S. mutans counts. Plates were incubated at 37ºC and 5% CO2, for 48h and the results were expressed by log10 scale of Colony-Forming Units (CFU/mL). Statistical Analyses Data were analyzed by means of the statistical software program SPSS version 20.0 (SPSS Inc., Chicago, USA). The Shapiro-Wilk test was used to verify the distribution of normality of the microbiologic results and of the percentage of hardness loss of the bovine tooth enamel blocks. With regard to the microbial count and the hardness loss values, ANOVA followed by Tukey test was used to verify whether there was statistical difference among the groups of treatment. Sample power was calculated between the groups EEP (tested substance) and the Chlorhexidine (positive control) using an α error of 5%, which resulted in a power of 0.8752.
Results The Hydroxycinnamic acids and derivatives content of the EEP at 33.33%, as well as the total phenolic compound content are expressed in Table 1. The pH of the EEP was 4.8 and the total solid content was 25.9 °Brix. The minimum inhibitory concentration of EEP against S. mutans was 2.08 mg/mL, and 8% for ethanol; while the minimum bactericide concentration of the extract and ethanol was 8.33mg/mL and 16%, respectively. After treating the biofilm formed on the bovine enamel fragments, G1 (EEP) percentage of hardness loss (%HL) was 84.41 ± 2.77, similar (p = 0.477) to G3 (Ethanol) (%HL = 87.80 ± 6.89). The %HL value (68.44 ± 12.98) of G2 (Chlorhexidine) was the lowest in comparison with the other groups (p = 0.010). G4 (Milli-Q water) presented the highest mean value for %HL (90.49±5.38) after the period of treatment. The groups G5 and G6, treated with EEP and Milli-Q water, respectively, but with no microorganisms, presented the following %HL: 16.11% ± 7.92; and 20.55% ± 10.65; respectively (p=0.952). All the results with reference to hardness loss of fragments may be visualized in Table 2. There was difference between G1 (EEP) and G4 (Milli-Q water) as regards the
8 number of S. mutans: 7.26±0.08 and 8.29±0.17CFU/mL, respectively (p = 0.001). The lower quantity of viable cells was presented by G2 (Chlorhexidine) (6.79±0.10 CFU/mL) (p = 0.043), and there was no difference (p = 0.435) between G3 and G4 (Table 2).
Discussion There are many strategies that can be implemented to control dental caries, such as: a low sugar diet, instruction to oral hygiene, use of fluoride dentifrice, the control of biofilm, and others. Besides that, there are evidences (Chandra Shekar, Nagarajappa, Suma, & Thakur, 2015) that support the advantages of natural products as coadjutant factors to be used in specific cases of caries activity. According to Chandra-Shekar, Nagarajappa, Suma, & Thakur (2015), the use of herbal extracts for the control of oral/dental diseases is considered to be an interesting alternative to synthetic antimicrobials due to their lower negative impact and to overcome intrinsic (primary) resistance or secondary resistance to drugs during therapy. Therefore, in an effort to reduce the prevalence of caries, different natural products have been previously tested for anticariogenic effect (Cheng, Li, He, & Zhou, 2015; Almeida et al., 2012). It is important to emphasize that the anticariogenic effect of a product involves both inhibition of growth of the bacteria responsible for caries disease, and inhibition of the enamel demineralization process (Antonio et al., 2011); effects tested in the present study. It is known that there is not only one bacterial species involved in the carious process (Kidd & Fejerskov, 2004; Gross et al., 2012). However, although Streptococcus mutans is not the only microorganism responsible for the onset of the disease, the biofilm formed by these species is usually chosen to evaluate the antibacterial and/or anticariogenic activities of different products (Bueno-Silva et al., 2013; Fernández, Giacaman, Tenuta, & Cury, 2015). According to Bueno-Silva et al. (2013), Streptococcus mutans and sucrose are key modulators of the development of cariogenic biofilms. Considering the antibacterial effect of phenolic compounds, studies have pointed out that these substances are the main active ingredients of natural products responsible for this function. They inhibit the enzyme glycosyltransferase, responsible among other factors, for the adherence of S. mutans to the tooth surface for dental biofilm formation (Koo et al., 2002). In the present study, the total polyphenol and hydroxycinnamic acids contents found in propolis was low (Table 1) in comparison with data from previous studies (Souza et al., 2007; Cabral et al., 2009). In spite of this finding, it is probable that the presence of caffeic and 5-caffeoylquinic acids (5-CQA) found in the tested extract, in synergy with other possible compounds, may have
9 contributed to decrease the number of viable microorganisms observed after treatment of the blocks/biofilm with EEP. According to Antonio et al. (2010), 5-CQA and caffeic acid are substances capable of inhibiting the growth of S. mutans. The microorganisms count in the group treated with EEP was significantly lower than that of the groups treated with 8% ethanol and Milli-Q water (Table 2). This result shows that EEP was efficient in reducing the number of S. mutans present in dental biofilm. However, the mean of the hardness loss percentage in the group (G1) treated with EEP (%HL = 84.41% ± 2.77) did not differ statistically from that of the groups treated with Milli-Q water (G4) and Ethanol (G3). According to Zhang et al. (2009), the enamel demineralization process may be inhibited by the interaction of polyphenols with the organic matrix, which results in lower mineral loss. These interactions involve covalent, ionic, and hydrogen bonding or hydrophobic processes, which induce the metamorphism of the enamel organic matrix. The metamorphic organic matrix is precipitated on the enamel, resulting in slowing down the speed of mineral ion loss, and consequently, enamel demineralization is inhibited. We did not find some of the phenolic components investigated (Table 1) and, consequently, a low variety of polyphenols of the propolis tested was observed. It possibly explains the low level of influence of this product in inhibiting the demineralization of the enamel analyzed, in spite of its capacity to act directly on the bacteria in conjunction with the other compounds, but the presence of other types of active polyphenols such as flavonoids, which were not individually analyzed in this study, is possible. This controversial result leads to the believe that the time of treatment may have played a fundamental role in this case, because lysis of the bacteria may have occurred in a period after the loss of hardness already in progress in the enamel; and the acidic pH (4.8) of the propolis extract may also have contributed to these results. Furthermore, no tests were performed with mixed biofilm and the authors suggest further studies on this subject. Also, although the power of the study presented a favorable result between the EEP and Chlorhexidine groups, the authors believe that a large sample size is needed in future studies. Another point to consider is the high concentration (80%) of ethanol that was used for biofilm treatment, which may have contributed to a smaller number of bacteria acting on dental demineralization, and therefore, to the results of hardness loss in G3 which, although higher, did not differ from those of G1. The minimum inhibitory concentration (MIC) of the substance was 2.08 mg/mL and the bactericidal concentration (MBC), 8.33 mg/mL. The values found are higher than those of previous studies (Cabral et al., 2009; Castro et al., 2009; Swerts, Costa, & Fiorini, 2010). The chemical composition of propolis is variable, according to its type,
10 origin and seasonality, which leads to its diverse biologic effects, such as antiinflammatory, antimicrobial and antioxidant, as shown in previous studies (Akao et al., 2003; Cabral et al., 2009). Therefore, this fact perfectly justifies the continuous study of propolis. For a better qualitative study of the extract as regards to its consumption, cytotoxicity tests must be conducted, which represents a limitation of the present study. However, in accordance with the administrative ruling of the Ministry of Agriculture of Brazil (Brasil, 2015), the technical standard for determination of identity and quality of propolis for consumption reads that the minimum content of phenolic compounds must be 0.50%. Thus, the contents found in the extract analyzed in the present study were above this value. Moreover, the cariogenic challenge with sucrose concentrations was constant up to the end of the experimental protocol, which also represents a limitation of this study. However, according to Pierro et al. (2014) a sudden exposure to an excess amount of sugar in laboratory studies may cause bacterial death because the rapid entry and degradation of sugars in cells can result in an accumulation of toxic levels of glycolytic intermediates. Thus, taking into consideration that it was an in vitro study, the authors only opted to frequently change the media to maintain the viability of the microorganisms. A previous study (Duarte et al., 2006) investigated the ethanolic extract of a novel type of propolis and its purified hexane fraction on S. mutans biofilms and the development of dental caries in rats. The results suggested that the cariostatic properties of propolis were due to the reduction of acid production and acid tolerance of cariogenic streptococci (Duarte et al., 2006). Recent clinical studies (Mohsin, Manohar, Rajesh, & Asif, 2015; Prabhakar, Karuna, Yavagal, & Deepak, 2015) reinforced the role of propolis, in their extract form or incorporated in dentifrices, to reducethe microbial load throughout the human oral cavity. Also, with the results obtained in the present research, the authors suggest that this propolis extract is a potent inhibitor of S. mutans biofilm but it was not able to inhibit the demineralization process of the dental enamel. However, in the future, comparison of the anticariogenic activity of different types of propolis, may possibly predict the most indicated types for use in dentistry.
Acknowledgments The authors thank the Fundação de Amparo à Pesquisa do Estado do Rio de Janeiro (FAPERJ) for the financial support and Sérgio Lutz Barbosa for the green própolis sample.
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15 Table 1: Hydroxycinnamic acids and derivatives presented in the ethanol extract of propolis Hydroxycinnamic acids and derivatives
(µg/mL)
3-caffeoylquinic acid
0.0
4-caffeoylquinic acid
4.84
5-caffeoylquinic acid
14.6
Caffeic acid
23.5
Ferulic acid
0.0
p-coumaric acid
0.0
3,4-dicaffeoylquinic acid
27.3
3,5-dicaffeoylquinic acid
18.4
4,5-dicaffeoylquinic acid
0.0
Total
88.64
16 Table 2: Percentage hardness loss (%) and number of Streptococcus mutans colonies present in biofilm after treatment of different studied groups (G) S. mutans (CFU/biofilm)
Group
Hardness loss (%)
G1
84.41±2.77a
7.26±0.08
G2
68.44±12.98b
6.79±0.10
G3
87.80±6.89a
8.00±0.05
G4
90.49±5.38a
8.29±0.17
a
b
c
c
Note: Study Groups: G1 – 33.33% EEP; G2 – 0.12% Chlorhexidine; G3 – 80% Ethanol c/v EEP; G4 – Milli-Q water. Different letters in the same column represent statistically significant results. Results of the microbial count are shown in log10 scale for better comparison.
