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In vitro antimicrobial activity of Caesalpinia ferrea Martius fruits against oral pathogens
Journal of Ethnopharmacology 124 (2009) 289–294
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In vitro antimicrobial activity of Caesalpinia ferrea Martius fruits against oral pathogens Fábio C. Sampaio a,e,∗ , Maria do Socorro V. Pereira b , Celidarque S. Dias c , Vicente Carlos O. Costa c , Nikeila C.O. Conde d , Marília A.R. Buzalaf e a
Department of Clinical and Social Dentistry, Brazil Department of Molecular Biology, Brazil Laboratory of Pharmaceutical Technology at the Federal University of Paraíba, 58051-900 João Pessoa, Brazil d Department of Oral Medicine, Federal University of Amazonas, 69033-760 Manaus, Brazil e Department of Biochemistry, São Paulo University, 17012-901 Bauru, Brazil b c
a r t i c l e
i n f o
Article history: Received 17 November 2008 Received in revised form 9 April 2009 Accepted 19 April 2009 Available online 3 May 2009 Keywords: Caesalpinia ferrea Martius Antimicrobial Oral biofilm
a b s t r a c t Aim: In the Amazon region of Brazil, the fruits of Caesalpinia ferrea Martius (Brazilian ironwood) are widely used as an antimicrobial and healing medicine in many situations including oral infections. This study aimed to evaluate the antimicrobial activity of Caesalpinia ferrea Martius fruit extract against oral pathogens. Materials and methods: Polyphenols estimation and spectral analysis (1 H NMR) of the methanol extract were carried out. The microorganisms Candida albicans, Streptococcus mutans, Streptococcus salivarius, Streptococcus oralis and Lactobacillus casei were tested using the microdilution method for planktonic cells (MIC) and a multispecies biofilm model. Chlorhexidine was used as positive control. Results: Polyphenols in the extract were estimated at 7.3% and 1 H NMR analysis revealed hydroxy phenols and methoxilated compounds. MIC values for Candida albicans, Streptococcus mutans, Streptococcus salivarius, Streptococcus oralis and Lactobacillus casei were 25.0, 40.0, 66.0, 100.0, 66.0 g/mL, respectively. For the biofilm assay, chlorhexidine and plant extract showed no growth at 10−4 and 10−5 microbial dilution, respectively. At 10−4 and 10−5 the growth values (mean ± SD) of the negative controls (DMSO and saline solution) for Streptococcus mutans, Streptococcus sp. and Candida albicans were 8.1 ± 0.7, 7.0 ± 0.6 and 5.9 ± 0.9 × 106 CFU, respectively. Conclusion: Caesalpinia ferrea fruit extract can inhibit in vitro growth of oral pathogens in planktonic and biofilm models supporting its use for oral infections. © 2009 Elsevier Ireland Ltd. All rights reserved.
1. Introduction Medicinal plants constitute the basis of health care systems in many Amazonian communities in Brazil. Ethnopharmacological studies in this region indicate Caesalpinia ferrea Martius (Brazilian ironwood) as an antifungal, antimicrobial and anti-inflammatory healing plant of the Amazonian forest. Locals use it in the form of tea (leaves, fruits or peel), syrup (peel) and as oral mouthwash steeping the fruits in alcohol for days (Vieira, 1992; Di Stasi and Hiruma-Lima, 2002; Borrás, 2003). The alcoholic tincture can be prepared in many ways such as placing the powder of the dried fruits (100–300 g) in alcohol (70%) for 8 days (Lorenzi and Matos,
∗ Corresponding author at: Department of Biological Sciences, Bauru Dental School, University of São Paulo, Al. Octávio Pinheiro Brisolla, 9-75, 17012-901 Bauru, SP, Brazil. Tel.: +55 14 32358346. E-mail address: [email protected] (F.C. Sampaio). 0378-8741/$ – see front matter © 2009 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.jep.2009.04.034
2002). A healing medicine can be also prepared with fresh fruits (500 g) and 100 mL of pure alcohol. The infusion of the fruits is recommended for healing oral wounds and controlling gastric problems (Cavalcante, 2008). In spite of its antimicrobial properties, most of the studies with the crude extract of Caesalpinia ferrea Martius had focused on its high content of polyphenols and the analgesic, anti-inflammatory, antiulcer and cancer chemopreventive properties (Bacchi et al., 1995; Queiroz et al., 2001; Nakamura et al., 2002). The crude extract of Caesalpinia ferrea Martius contains anthraquinones, alkaloids, depsides, depsidones, flavonoids, lactones, saponins, sugars, tannins, sesquiterpenes and triterpenes. Tannins are regarded as the major component (Souza et al., 2006). Isolated compounds from Brazilian ironwood fruits can inhibit aldose reductase suggesting an anti-diabetes effect (Ueda et al., 2001). Plants of the same genus (Caesalpinia L.) that are used for sore throat and oral infections have antibacterial properties against both Gram-positive and Gram-negative bacteria
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(Carvalho et al., 1996; Saeed and Sabir, 2001; Sudhakar et al., 2006). It has been suggested that polyphenols are responsible for the antibacterial properties upon Streptococcus mutans (Kakiuchi et al., 1986). This can be related to polyphenols action as mediators in cell membrane and oxidative phosphorylation at low concentrations (Scalbert et al., 2005). However, only recently, polyphenols in fruits and leaves have received attention regarding their antibacterial effect upon microorganisms in biofilms (Huber et al., 2003; Duarte et al., 2006; Percival et al., 2006). Hence, the aim of this study was to evaluate the antimicrobial activity of the crude extract of Caesalpinia ferrea Martius (Brazilian ironwood) fruits against oral pathogens organized as planktonic cells and also in a biofilm model. 2. Materials and methods 2.1. Plant material Caesalpinia ferrea Martius fruits. 2.1.1. Source, collection and identification The fruits from Caesalpinia ferrea Martius were collected at the experimental farm of Federal University of Amazonas in Manaus (Brazil) in December 2005 with authorization for collection and transportation from IBAMA (Brazilian Institute of the Environment and Natural Renewable Resources) number 044/2004. A voucher was deposited at Lauro Pires Xavier Herbarium at the Federal University of Paraíba, João Pessoa, PB, Brazil (JPB-38756). 2.1.2. Extraction The fruits (200 g) were dried at room temperature and grounded prior preparation of the extract. The extraction was carried out by using methanol (80%, v/v) for a period of 72 h without any heating procedure. The final volume of the filtrate was reduced by rotaryevaporation (Fisatom 802, Sao Paulo, Brazil) to yield 48.4 g (24%, w/w) and reaching a 0.5 g/mL solution. 2.2. Quantification and characterization of polyphenols 2.2.1. Prussian Blue Method Total phenolics in the crude extract were estimated by the Prussian Blue Method as described previously (Graham, 1992). All solutions were prepared from analytical grade reagents and deionized water was used throughout. Solution 1 was prepared with 0.02 M ferric chloride diluted in 0.10 M HCl; solution 2 was 0.016 M of potassium ferricyanide and solution 3 (stabilizer) was prepared with 1% gum arabic, 10 mL of 85% phosphoric acid and 30 mL of deionized water. Standards (in triplicate) were 0.001 and 0.01 M gallic acid monohydrate (3,4,5-trihydroxybenzoic acid) dissolved in methanol. A blank solution was prepared with methanol and 3 mL of deionized water. Samples were prepared dispensing 100 L or smaller appropriate volumes made up to 100 L with solvent (methanol) into a test tube. A set of three samples was finally diluted in 3 mL of deionized water following the guidelines. To each sample and standard solution 2 was added followed by solution 1. The mixture went to a vortex for 30 s and after 15 min, solution 3 was added. The absorbance was read at 700 nm in a spectrophotometer (Beckman DU640, Beckman, Fullerton, CA, USA) within the appropriate timeline, just after color characterization stability. 2.2.2. Spectral analysis Spectral analysis of the extract was performed for general basic phenol characterization. The analysis was carried out on a Mercury-Varian 200 MHz (1 H) nuclear magnetic resonance (NMR) spectrometer optimized for 1D and 2D techniques. 10 mg of the
extract diluted in 0.6 mL of deutered dimethyl sulfoxide (DMSOD6 ), recorded from typical peaks of the nondeutered portion of the NMR solvent in RMN 1 H in relation to TMS, was used. 2.3. Microorganisms and biofilm assays 2.3.1. Microorganisms The following microorganisms were used in the microbiological assays: Candida albicans (ATCC 36232), Streptococcus mutans (ATCC 25175), Streptococcus salivarius (ATCC 7073), Streptococcus oralis (ATCC 10557) and Lactobacillus casei (ATCC 7469). The bacterial strains were cultured in brain heart infusion (BHI) (Difco, MI, USA) and for Candida albicans Sabouraud-dextrose broth (Difco, MI, USA) was used. The microorganisms were incubated under aerobic conditions at 37 ◦ C. 2.3.2. Microdilution method The Minimum Inhibitory Concentration (MIC) values of the crude extract were determined using the resazurin microdilution broth assay for microplates (Andrews, 2001; Sarker et al., 2007), and NCCLS method for yeast (NCCLS, 2002) with few modifications. Briefly, the crude extract was dissolved in DMSO (1 mg/mL) and then diluted in BHI broth (Difco, MI, USA) to achieve concentrations ranging from 400 to 10 g/mL. For MIC values below 10 g/mL, all procedures were repeated with final values ranging from 10.0 to 0.3 g/mL. Saline solution and DMSO 10% (v/v) were used as negative controls. The inocula were adjusted to each microorganism to yield a cell concentration of 108 CFU/mL. A final volume of 100 L was achieved in each well. One well with specific medium and microorganism was used as control of the growth, and one inoculated well was free of the test extract to check the sterility of the media. The microplates were prepared in triplicate and incubated at 37 ◦ C for 24 h. A volume of 30 L of a homogenous aqueous solution of resazurin (0.01%) was added to each well to indicate viability of bacteria and yeast (Sarker et al., 2007). The MIC endpoint was identified as the lowest concentration at which there was no visible growth indicated by the resazurin original blue color. 2.3.3. Multispecies biofilm model The antimicrobial activity of the extract was also evaluated using a multispecies biofilm model (Guggenheim et al., 2004). All microorganisms were cultivated independently in fluid thioglycolate medium (Difco Laboratories, Detroit, USA) supplemented with 67 mmol/L of Sorensen’s solution, pH 7.2 named Universal Medium (UM). After grown in UM, 200 L of each microorganism suspension were transferred for UM–glucose (1× 5 mL Sorensen’s solution + 0.3% de glucose) and incubated for 7 h. The suspensions were diluted (10−4 ) and 1 mL of each diluted suspension was combined and placed at cold temperature for 6 h. Approximately 500 mL of unstimulated saliva were collected from three volunteers for 1 h during several days always 2 h after eating, drinking or brushing their teeth. The saliva was centrifuged (30 min, 4 ◦ C, 20,000 × g) and the supernatant was vacuum-filtered in a very low protein binding filter system (StericupTM 0.22 m, Millipore Corporation, Billerica, MA, USA). Absence of any viable microorganism in the saliva was tested by incubating the material in Mitis Salivarius Bacitracin Agar (Difco Laboratories, Detroit, USA), Muller Hinton Agar (Difco Laboratories, Detroit, USA) and Sabouraud Agar (Difco Laboratories, Detroit, USA). The biofilm was formed on blocks (n = 6) of sterilized bovine teeth (10.6 mm diameter × 5 mm height) standardized by size and weight in a saliva:broth medium (ratio of 60:40) for 3 consecutive days. In order to form acquired pellicle, each block was incubated in saliva for 4 h at room temperature. Then the saliva was taken, and the test tubes were filled with a mixture of saliva (960 L) + UM (640 L) with 0.3% glucose plus a pool of microorganisms (200 L). The blocks were incubated for
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Fig. 1. Schematic representation of the multispecies biofilm model. The groups of solutions in bath 1 and 2 were: plant extract, saline solution or DMSO (negative controls), chlorhexidine (positive control).
64 h and 30 min in polypropylene test tubes. Medium was renovated after 16 h and 30 min of incubation and later at intervals of 24 h until the last day (Fig. 1). Saline solution and chlorhexidine gluconate (0.12%, Periogard, Colgate-Palmolive, Brazil) were used as negative and positive controls, respectively. The biofilms were exposed to controls and test solutions (Caesalpinia ferrea Martius extract suspended in deionized water 1:1) in a separate test tube for 1 min on two occasions: at 16 h and 30 min, and at 40 h and 30 min of incubation. After each solution bath, the blocks were carefully immersed in saline solutions (3× 2 mL) for 10 s. All procedures were in duplicate and the biofilms were also dipped (for 10 s) three times a day in saline solution for passages through an air–liquid interface (Fig. 1). After 64 h and 30 min of incubation, the enamel blocks were shaken in saline solution (1 mL) and sonicated for 30 min. Serial dilutions of sonicated cells (100 to 10−6 ) were prepared and cultivated in Mitis Salivarius Bacitracin agar (Difco Laboratories, Detroit, USA) and Mueller Hinton agar (Difco Laboratories, Detroit, USA) (37 ◦ C, micro-aerophilic incubation), and in Sabouraud dextrose agar (Difco Laboratories, Detroit, USA) (37 ◦ C, aerophilic incubation) for final CFU counting.
the presence of sugar protons and aliphatic H atoms. Furthermore, at least three CH3 groups can be observed as singlets at ı 3.57 (OCH3 ), 3.31 and 3.15 ppm (Fig. 4). Minimal Inhibitory Concentration values for Candida albicans, Streptococcus mutans, Streptococcus salivarius, Streptococcus oralis and Lactobacillus casei were 25.0, 40.0, 66.0, 100.0, 66.0 g/mL, respectively. For these microorganisms chlorhexidine values were 15.0, 0.4, 0.4, 1.5, 0.8 g/mL. At >10−3 microbial dilution both products showed cell growth. At 10−4 microbial dilution, chlorhexidine showed no growth. Taking this cell dilution as reference, the results for the extract upon Streptococcus mutans, Streptococcus sp. and Candida albicans were 0.5 ± 0.1, 0.5 ± 0.0 and 0.7 ± 0.1 × 106 CFU, respectively. For negative controls, the results (10−4 and 10−5 ) were 8.1 ± 0.7, 7.0 ± 0.6 and 5.9 ± 0.9 × 106 CFU. No growth was observed at 10−5 for both compounds. The absence of cell growth in 10−5 microbial dilution indicates that the plant extract has antimicrobial activity on biofilm formation. However, this activity is lower when compared to chlorhexidine. Hence, these findings suggest that the resistance of the microbial biofilm to the plant extract starts at lower bacterial biofilm population when compared to the gold standard.
3. Results 4. Discussion The amount of polyphenols in the crude hydromethanolic extract of Caesalpinia ferrea Martius was estimated at 7.3%. The 1 H NMR spectrum of the extract presents characteristic signals of aromatic protons in the ı 7.46–7.48 as well as 6.90–7.08 and 6.38–6.58 ppm range (Figs. 2 and 3). In addition, other overlapping signals can be seen in the ı 6.14–6.25 and 2.9–5.5 region, suggesting
Many microbiological studies tested oral antimicrobials growing cells in a planktonic state (Marsh, 2004; Scheie and Petersen, 2004). This is also valid for Brazilian medicinal plants when evaluated as oral antimicrobials (Vasconcelos et al., 2006). The understanding of oral ecological imbalances in microbial biofilms are of great
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Fig. 2.
1
H NMR spectrum of the extract of Caesalpinia ferrea Martius fruits.
importance to develop strategies to prevent dental caries and periodontal diseases since oral microorganisms organized in biofilms are more resistant to immune defense mechanisms and less susceptible to chemical agents (Davies, 2003; Marsh, 2006; Song et al., 2007; Quave et al., 2008). Therefore, the MIC reference values (end points) are useful, but these data must be taken with caution when considering biofilm related oral diseases. The advantage of the in vitro multispecies Zürich biofilm model is the possibility to mimic the oral environment (e.g. saliva, acquired
Fig. 3.
1
pellicle, tooth surface, antimicrobial challenge) without the need of expensive laboratory equipment. In our study, few modifications in the original model were performed (Guggenheim et al., 2001). For instance, the bacterial selection was virtually the same and the saliva:broth ratio of 60:40 proved to be better in our model than the 70:30 original ratio for the growing pool of bacteria. Sterilizing saliva by filtration provided better growth than pasteurized saliva probably because protein denaturation was avoided. Rich protein saliva is important to promote cell adhesion to tooth surface and
H NMR spectrum in the 6.0–7.6 ppm range.
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Fig. 4.
1
293
H NMR spectrum in the 1.0–5.5 ppm range.
the protein–polyphenols complexation in this model (Guggenheim et al., 2001; Spencer et al., 1988; Bennick, 2002). In order to compensate any loss in salivary proteins due to filtration, strains of Streptococcus salivarius was included in our biofilm model (Williams and Kraus, 1963). The Streptococcus salivarius is not an oral pathogen in mature dental biofilms, but it is ubiquitous in saliva and can produce large amounts of extracellular polymers when exposed to sucrose. This bacterium is a pioneer colonizing in dental plaque and creates favorable conditions so that other species can begin colonization (Marsh, 2005). Our biofilm model reflected an immature dental biofilm since we did not include Fusobacterium nucleatum, an oral bacterium related to bacteria co-aggregation and periodontal diseases. Nevertheless, an oral biofilm was created and the environmental conditions in our biofilm model were suitable for bacteria growth as shown by the data of the negative controls. Our results showed that the extract of Caesalpinia ferrea Martius has a good inhibitory effect upon oral microorganism in a planktonic state since MIC values ranged between 25 and 100 g/mL. A less pronounced effect was observed upon bacteria in a biofilm form since viable bacteria at 10−4 microbial dilution could be found. This last result was expected since bacterial biofilms are several hundred times more resistant than the MIC for the same bacteria in planktonic state. Our data suggest that the Amazonian tradition of using this plant for oral hygiene or infections is effective. Moreover, it can be speculated that the crude extract of this plant is active against free salivary oral bacteria in vivo. The biofilm model was a valuable attempt to simulate the plant mouthwash under in vivo conditions. Our findings support that the extract was effective but with lower activity when compared to a regular chlorhexidine mouthwash (1.2 mg/mL). However, for this experiment the extract was diluted in water in order to avoid interference in cell adhesion due to the high polarity of DMSO (Marshall et al., 1989). Thus, the plant extract in the biofilm assay was basically composed of water-soluble compounds and some biological activity could be compromised in this process. The 1 H NMR spectrum
confirms the presence of several chemical constituents including polyphenols and many other organic compounds (Cowan, 1999; Souza et al., 2006). The evident presence of tannins in the extract can certainly provoke salivary protein biding affecting salivary bacteria, but other active constituents in the extract are certainly contributing for this biological activity and other antimicrobial mechanisms of action. For instance, aiming to control dental caries, some alkaloids, flavonoids and tannins have recently proved to be effective against the same cariogenic bacteria species evaluated in our study (Badria and Zidan, 2004). Under clinical conditions the antibacterial effect of many natural products might be reduced due to constant salivary flow whereas chlorhexidine is less affected by saliva due to its high substantivity (Scheie and Petersen, 2004). Except for essential oils few natural compounds were tested in vivo for their ability to be present in the oral cavity for longer periods. Tea polyphenols molecules can be adsorbed and affect the acquired salivary pellicle (Joiner et al., 2004). This is an indication that polyphenols may interfere more on biofilm formation than on cells of a mature biofilm. In our study, a pre-exposure of the blocks to the plant extract was not tested and this effect could not be evaluated. This seems to be true since cocoa polyphenols reduces pH production without affecting Streptococcus mutans growth and that the crude extract of cranberry can affect the accumulation and polysaccharide composition of Streptococcus mutans biofilms (Duarte et al., 2006; Percival et al., 2006). The synergic or isolated activity of other constituents was not evaluated in our study, but this approach may render interesting results for controlling oral biofilm growth. For prophylactic purposes, many authors support that the active agents of antimicrobials should prevent biofilm formation without affecting the biological equilibrium within the oral cavity (Baehni and Takeuchi, 2003; Scheie and Petersen, 2004; Marsh and Percival, 2006). Since this is a characteristic of many organic compounds, medicine plants can certainly be a valuable source of oral antimicrobials (Cowan, 1999). This is particularly important when active
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compounds can be found in the fruits of plants such as Brazilian ironwood which are available in many local markets of the Amazonian region. Finally, our findings support that the fruits of Caesalpinia ferrea Martius have antibacterial activity against oral pathogens in planktonic or biofilm in vitro models. Acknowledgements This study was supported by CNPq 306234/2004-1 and FAPEAM—PROGRAMA POSVINC 1192-4-A. References Andrews, J.M., 2001. Determination of minimum inhibitory concentrations. Journal of Antimicrobial Chemotherapy 48 (Suppl. 1), 5–16. Bacchi, E.M., Sertie, J.A., Villa, N., Katz, H., 1995. Antiulcer action and toxicity of Styrax camporum and Caesalpinia ferrea. Planta Medica 61, 204–207. Badria, F.A., Zidan, O.A., 2004. Natural products for dental caries prevention. Journal of Medicinal Food 7, 381–384. Baehni, P.C., Takeuchi, Y., 2003. Anti-plaque agents in the prevention of biofilmassociated oral diseases. Oral Diseases 9 (Suppl. 1), 23–29. Bennick, A., 2002. Interaction of plant polyphenols with salivary proteins. Critical Reviews in Oral Biology and Medicine 13, 184–196. Borrás, M.R.L., 2003. Plantas da Amazônia: Medicinais ou mágica?—Plantas comercializadas no mercado Adolpho Lisboa. Editora Valer/Governo do Estado do Amazonas, Manaus. Carvalho, J.C.T., Teixeira, J.R.M., Souza, P.J.C., Bastos, J.K., dos Santos Filho, D., Sarti, S.J., 1996. Preliminary studies of analgesic and anti-inflammatory properties of Caesalpinia ferrea crude extract. Journal of Ethnopharmacology 53, 175–178. Cavalcante, R., 2008. As plantas medicinais na Odontologia: um guia prático. Expressão Gráfica, Rio Branco. Cowan, M.M., 1999. Plant products as antimicrobial agents. Clinical Microbiology Reviews 12, 564–582. Davies, D., 2003. Understanding biofilm resistance to antibacterial agents. Nature Reviews Drug Discovery 2, 114–122. Di Stasi, L.C., Hiruma-Lima, C.A., 2002. Plantas medicinais na Amazônia e na Mata Atlântica. Unesp, São Paulo. Duarte, S., Gregoire, S., Singh, A.P., Vorsa, N., Schaich, K., Bowen, W.H., Koo, H., 2006. Inhibitory effects of cranberry polyphenols on formation and acidogenicity of Streptococcus mutans biofilms. FEMS Microbiology Letters 257, 50–56. Graham, H.D., 1992. Stabilization of Prussian blue color in the determination of polyphenols. Journal of Agricultural and Food Chemistry 40, 801–805. Guggenheim, B., Giertsen, E., Schupbach, P., Shapiro, S., 2001. Validation of an in vitro biofilm model of supragingival plaque. Journal of Dental Research 80, 363–370. Guggenheim, B., Guggenheim, M., Gmur, R., Giertsen, E., Thurnheer, T., 2004. Application of the Zurich biofilm model to problems of cariology. Caries Research 38, 212–222. Huber, B., Eberl, L., Feucht, W., Polster, J., 2003. Influence of polyphenols on bacterial biofilm formation and quorum-sensing. Zeitschrift für Naturforschung C 58, 879–884. Joiner, A., Muller, D., Elofsson, U.M., Arnebrant, T., 2004. Ellipsometry analysis of the in vitro adsorption of tea polyphenols onto salivary pellicles. European Journal of Oral Sciences 112, 510–515. Kakiuchi, N., Hattori, M., Nishizawa, M., Yamagishi, T., Okuda, T., Namba, T., 1986. Studies on dental caries prevention by traditional medicines. VIII. Inhibitory effect of various tannins on glucan synthesis by glucosyltransferase from Streptococcus mutans. Chemical and Pharmaceutical Bulletin (Tokyo) 34, 720–725. Lorenzi, H., Matos, F.J.A., 2002. Plantas Medicinais no Brasil. Instituto Plantarum, Nova Odessa.
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Journal of Ethnopharmacology journal homepage: www.elsevier.com/locate/jethpharm
In vitro antimicrobial activity of Caesalpinia ferrea Martius fruits against oral pathogens Fábio C. Sampaio a,e,∗ , Maria do Socorro V. Pereira b , Celidarque S. Dias c , Vicente Carlos O. Costa c , Nikeila C.O. Conde d , Marília A.R. Buzalaf e a
Department of Clinical and Social Dentistry, Brazil Department of Molecular Biology, Brazil Laboratory of Pharmaceutical Technology at the Federal University of Paraíba, 58051-900 João Pessoa, Brazil d Department of Oral Medicine, Federal University of Amazonas, 69033-760 Manaus, Brazil e Department of Biochemistry, São Paulo University, 17012-901 Bauru, Brazil b c
a r t i c l e
i n f o
Article history: Received 17 November 2008 Received in revised form 9 April 2009 Accepted 19 April 2009 Available online 3 May 2009 Keywords: Caesalpinia ferrea Martius Antimicrobial Oral biofilm
a b s t r a c t Aim: In the Amazon region of Brazil, the fruits of Caesalpinia ferrea Martius (Brazilian ironwood) are widely used as an antimicrobial and healing medicine in many situations including oral infections. This study aimed to evaluate the antimicrobial activity of Caesalpinia ferrea Martius fruit extract against oral pathogens. Materials and methods: Polyphenols estimation and spectral analysis (1 H NMR) of the methanol extract were carried out. The microorganisms Candida albicans, Streptococcus mutans, Streptococcus salivarius, Streptococcus oralis and Lactobacillus casei were tested using the microdilution method for planktonic cells (MIC) and a multispecies biofilm model. Chlorhexidine was used as positive control. Results: Polyphenols in the extract were estimated at 7.3% and 1 H NMR analysis revealed hydroxy phenols and methoxilated compounds. MIC values for Candida albicans, Streptococcus mutans, Streptococcus salivarius, Streptococcus oralis and Lactobacillus casei were 25.0, 40.0, 66.0, 100.0, 66.0 g/mL, respectively. For the biofilm assay, chlorhexidine and plant extract showed no growth at 10−4 and 10−5 microbial dilution, respectively. At 10−4 and 10−5 the growth values (mean ± SD) of the negative controls (DMSO and saline solution) for Streptococcus mutans, Streptococcus sp. and Candida albicans were 8.1 ± 0.7, 7.0 ± 0.6 and 5.9 ± 0.9 × 106 CFU, respectively. Conclusion: Caesalpinia ferrea fruit extract can inhibit in vitro growth of oral pathogens in planktonic and biofilm models supporting its use for oral infections. © 2009 Elsevier Ireland Ltd. All rights reserved.
1. Introduction Medicinal plants constitute the basis of health care systems in many Amazonian communities in Brazil. Ethnopharmacological studies in this region indicate Caesalpinia ferrea Martius (Brazilian ironwood) as an antifungal, antimicrobial and anti-inflammatory healing plant of the Amazonian forest. Locals use it in the form of tea (leaves, fruits or peel), syrup (peel) and as oral mouthwash steeping the fruits in alcohol for days (Vieira, 1992; Di Stasi and Hiruma-Lima, 2002; Borrás, 2003). The alcoholic tincture can be prepared in many ways such as placing the powder of the dried fruits (100–300 g) in alcohol (70%) for 8 days (Lorenzi and Matos,
∗ Corresponding author at: Department of Biological Sciences, Bauru Dental School, University of São Paulo, Al. Octávio Pinheiro Brisolla, 9-75, 17012-901 Bauru, SP, Brazil. Tel.: +55 14 32358346. E-mail address: [email protected] (F.C. Sampaio). 0378-8741/$ – see front matter © 2009 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.jep.2009.04.034
2002). A healing medicine can be also prepared with fresh fruits (500 g) and 100 mL of pure alcohol. The infusion of the fruits is recommended for healing oral wounds and controlling gastric problems (Cavalcante, 2008). In spite of its antimicrobial properties, most of the studies with the crude extract of Caesalpinia ferrea Martius had focused on its high content of polyphenols and the analgesic, anti-inflammatory, antiulcer and cancer chemopreventive properties (Bacchi et al., 1995; Queiroz et al., 2001; Nakamura et al., 2002). The crude extract of Caesalpinia ferrea Martius contains anthraquinones, alkaloids, depsides, depsidones, flavonoids, lactones, saponins, sugars, tannins, sesquiterpenes and triterpenes. Tannins are regarded as the major component (Souza et al., 2006). Isolated compounds from Brazilian ironwood fruits can inhibit aldose reductase suggesting an anti-diabetes effect (Ueda et al., 2001). Plants of the same genus (Caesalpinia L.) that are used for sore throat and oral infections have antibacterial properties against both Gram-positive and Gram-negative bacteria
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(Carvalho et al., 1996; Saeed and Sabir, 2001; Sudhakar et al., 2006). It has been suggested that polyphenols are responsible for the antibacterial properties upon Streptococcus mutans (Kakiuchi et al., 1986). This can be related to polyphenols action as mediators in cell membrane and oxidative phosphorylation at low concentrations (Scalbert et al., 2005). However, only recently, polyphenols in fruits and leaves have received attention regarding their antibacterial effect upon microorganisms in biofilms (Huber et al., 2003; Duarte et al., 2006; Percival et al., 2006). Hence, the aim of this study was to evaluate the antimicrobial activity of the crude extract of Caesalpinia ferrea Martius (Brazilian ironwood) fruits against oral pathogens organized as planktonic cells and also in a biofilm model. 2. Materials and methods 2.1. Plant material Caesalpinia ferrea Martius fruits. 2.1.1. Source, collection and identification The fruits from Caesalpinia ferrea Martius were collected at the experimental farm of Federal University of Amazonas in Manaus (Brazil) in December 2005 with authorization for collection and transportation from IBAMA (Brazilian Institute of the Environment and Natural Renewable Resources) number 044/2004. A voucher was deposited at Lauro Pires Xavier Herbarium at the Federal University of Paraíba, João Pessoa, PB, Brazil (JPB-38756). 2.1.2. Extraction The fruits (200 g) were dried at room temperature and grounded prior preparation of the extract. The extraction was carried out by using methanol (80%, v/v) for a period of 72 h without any heating procedure. The final volume of the filtrate was reduced by rotaryevaporation (Fisatom 802, Sao Paulo, Brazil) to yield 48.4 g (24%, w/w) and reaching a 0.5 g/mL solution. 2.2. Quantification and characterization of polyphenols 2.2.1. Prussian Blue Method Total phenolics in the crude extract were estimated by the Prussian Blue Method as described previously (Graham, 1992). All solutions were prepared from analytical grade reagents and deionized water was used throughout. Solution 1 was prepared with 0.02 M ferric chloride diluted in 0.10 M HCl; solution 2 was 0.016 M of potassium ferricyanide and solution 3 (stabilizer) was prepared with 1% gum arabic, 10 mL of 85% phosphoric acid and 30 mL of deionized water. Standards (in triplicate) were 0.001 and 0.01 M gallic acid monohydrate (3,4,5-trihydroxybenzoic acid) dissolved in methanol. A blank solution was prepared with methanol and 3 mL of deionized water. Samples were prepared dispensing 100 L or smaller appropriate volumes made up to 100 L with solvent (methanol) into a test tube. A set of three samples was finally diluted in 3 mL of deionized water following the guidelines. To each sample and standard solution 2 was added followed by solution 1. The mixture went to a vortex for 30 s and after 15 min, solution 3 was added. The absorbance was read at 700 nm in a spectrophotometer (Beckman DU640, Beckman, Fullerton, CA, USA) within the appropriate timeline, just after color characterization stability. 2.2.2. Spectral analysis Spectral analysis of the extract was performed for general basic phenol characterization. The analysis was carried out on a Mercury-Varian 200 MHz (1 H) nuclear magnetic resonance (NMR) spectrometer optimized for 1D and 2D techniques. 10 mg of the
extract diluted in 0.6 mL of deutered dimethyl sulfoxide (DMSOD6 ), recorded from typical peaks of the nondeutered portion of the NMR solvent in RMN 1 H in relation to TMS, was used. 2.3. Microorganisms and biofilm assays 2.3.1. Microorganisms The following microorganisms were used in the microbiological assays: Candida albicans (ATCC 36232), Streptococcus mutans (ATCC 25175), Streptococcus salivarius (ATCC 7073), Streptococcus oralis (ATCC 10557) and Lactobacillus casei (ATCC 7469). The bacterial strains were cultured in brain heart infusion (BHI) (Difco, MI, USA) and for Candida albicans Sabouraud-dextrose broth (Difco, MI, USA) was used. The microorganisms were incubated under aerobic conditions at 37 ◦ C. 2.3.2. Microdilution method The Minimum Inhibitory Concentration (MIC) values of the crude extract were determined using the resazurin microdilution broth assay for microplates (Andrews, 2001; Sarker et al., 2007), and NCCLS method for yeast (NCCLS, 2002) with few modifications. Briefly, the crude extract was dissolved in DMSO (1 mg/mL) and then diluted in BHI broth (Difco, MI, USA) to achieve concentrations ranging from 400 to 10 g/mL. For MIC values below 10 g/mL, all procedures were repeated with final values ranging from 10.0 to 0.3 g/mL. Saline solution and DMSO 10% (v/v) were used as negative controls. The inocula were adjusted to each microorganism to yield a cell concentration of 108 CFU/mL. A final volume of 100 L was achieved in each well. One well with specific medium and microorganism was used as control of the growth, and one inoculated well was free of the test extract to check the sterility of the media. The microplates were prepared in triplicate and incubated at 37 ◦ C for 24 h. A volume of 30 L of a homogenous aqueous solution of resazurin (0.01%) was added to each well to indicate viability of bacteria and yeast (Sarker et al., 2007). The MIC endpoint was identified as the lowest concentration at which there was no visible growth indicated by the resazurin original blue color. 2.3.3. Multispecies biofilm model The antimicrobial activity of the extract was also evaluated using a multispecies biofilm model (Guggenheim et al., 2004). All microorganisms were cultivated independently in fluid thioglycolate medium (Difco Laboratories, Detroit, USA) supplemented with 67 mmol/L of Sorensen’s solution, pH 7.2 named Universal Medium (UM). After grown in UM, 200 L of each microorganism suspension were transferred for UM–glucose (1× 5 mL Sorensen’s solution + 0.3% de glucose) and incubated for 7 h. The suspensions were diluted (10−4 ) and 1 mL of each diluted suspension was combined and placed at cold temperature for 6 h. Approximately 500 mL of unstimulated saliva were collected from three volunteers for 1 h during several days always 2 h after eating, drinking or brushing their teeth. The saliva was centrifuged (30 min, 4 ◦ C, 20,000 × g) and the supernatant was vacuum-filtered in a very low protein binding filter system (StericupTM 0.22 m, Millipore Corporation, Billerica, MA, USA). Absence of any viable microorganism in the saliva was tested by incubating the material in Mitis Salivarius Bacitracin Agar (Difco Laboratories, Detroit, USA), Muller Hinton Agar (Difco Laboratories, Detroit, USA) and Sabouraud Agar (Difco Laboratories, Detroit, USA). The biofilm was formed on blocks (n = 6) of sterilized bovine teeth (10.6 mm diameter × 5 mm height) standardized by size and weight in a saliva:broth medium (ratio of 60:40) for 3 consecutive days. In order to form acquired pellicle, each block was incubated in saliva for 4 h at room temperature. Then the saliva was taken, and the test tubes were filled with a mixture of saliva (960 L) + UM (640 L) with 0.3% glucose plus a pool of microorganisms (200 L). The blocks were incubated for
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Fig. 1. Schematic representation of the multispecies biofilm model. The groups of solutions in bath 1 and 2 were: plant extract, saline solution or DMSO (negative controls), chlorhexidine (positive control).
64 h and 30 min in polypropylene test tubes. Medium was renovated after 16 h and 30 min of incubation and later at intervals of 24 h until the last day (Fig. 1). Saline solution and chlorhexidine gluconate (0.12%, Periogard, Colgate-Palmolive, Brazil) were used as negative and positive controls, respectively. The biofilms were exposed to controls and test solutions (Caesalpinia ferrea Martius extract suspended in deionized water 1:1) in a separate test tube for 1 min on two occasions: at 16 h and 30 min, and at 40 h and 30 min of incubation. After each solution bath, the blocks were carefully immersed in saline solutions (3× 2 mL) for 10 s. All procedures were in duplicate and the biofilms were also dipped (for 10 s) three times a day in saline solution for passages through an air–liquid interface (Fig. 1). After 64 h and 30 min of incubation, the enamel blocks were shaken in saline solution (1 mL) and sonicated for 30 min. Serial dilutions of sonicated cells (100 to 10−6 ) were prepared and cultivated in Mitis Salivarius Bacitracin agar (Difco Laboratories, Detroit, USA) and Mueller Hinton agar (Difco Laboratories, Detroit, USA) (37 ◦ C, micro-aerophilic incubation), and in Sabouraud dextrose agar (Difco Laboratories, Detroit, USA) (37 ◦ C, aerophilic incubation) for final CFU counting.
the presence of sugar protons and aliphatic H atoms. Furthermore, at least three CH3 groups can be observed as singlets at ı 3.57 (OCH3 ), 3.31 and 3.15 ppm (Fig. 4). Minimal Inhibitory Concentration values for Candida albicans, Streptococcus mutans, Streptococcus salivarius, Streptococcus oralis and Lactobacillus casei were 25.0, 40.0, 66.0, 100.0, 66.0 g/mL, respectively. For these microorganisms chlorhexidine values were 15.0, 0.4, 0.4, 1.5, 0.8 g/mL. At >10−3 microbial dilution both products showed cell growth. At 10−4 microbial dilution, chlorhexidine showed no growth. Taking this cell dilution as reference, the results for the extract upon Streptococcus mutans, Streptococcus sp. and Candida albicans were 0.5 ± 0.1, 0.5 ± 0.0 and 0.7 ± 0.1 × 106 CFU, respectively. For negative controls, the results (10−4 and 10−5 ) were 8.1 ± 0.7, 7.0 ± 0.6 and 5.9 ± 0.9 × 106 CFU. No growth was observed at 10−5 for both compounds. The absence of cell growth in 10−5 microbial dilution indicates that the plant extract has antimicrobial activity on biofilm formation. However, this activity is lower when compared to chlorhexidine. Hence, these findings suggest that the resistance of the microbial biofilm to the plant extract starts at lower bacterial biofilm population when compared to the gold standard.
3. Results 4. Discussion The amount of polyphenols in the crude hydromethanolic extract of Caesalpinia ferrea Martius was estimated at 7.3%. The 1 H NMR spectrum of the extract presents characteristic signals of aromatic protons in the ı 7.46–7.48 as well as 6.90–7.08 and 6.38–6.58 ppm range (Figs. 2 and 3). In addition, other overlapping signals can be seen in the ı 6.14–6.25 and 2.9–5.5 region, suggesting
Many microbiological studies tested oral antimicrobials growing cells in a planktonic state (Marsh, 2004; Scheie and Petersen, 2004). This is also valid for Brazilian medicinal plants when evaluated as oral antimicrobials (Vasconcelos et al., 2006). The understanding of oral ecological imbalances in microbial biofilms are of great
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Fig. 2.
1
H NMR spectrum of the extract of Caesalpinia ferrea Martius fruits.
importance to develop strategies to prevent dental caries and periodontal diseases since oral microorganisms organized in biofilms are more resistant to immune defense mechanisms and less susceptible to chemical agents (Davies, 2003; Marsh, 2006; Song et al., 2007; Quave et al., 2008). Therefore, the MIC reference values (end points) are useful, but these data must be taken with caution when considering biofilm related oral diseases. The advantage of the in vitro multispecies Zürich biofilm model is the possibility to mimic the oral environment (e.g. saliva, acquired
Fig. 3.
1
pellicle, tooth surface, antimicrobial challenge) without the need of expensive laboratory equipment. In our study, few modifications in the original model were performed (Guggenheim et al., 2001). For instance, the bacterial selection was virtually the same and the saliva:broth ratio of 60:40 proved to be better in our model than the 70:30 original ratio for the growing pool of bacteria. Sterilizing saliva by filtration provided better growth than pasteurized saliva probably because protein denaturation was avoided. Rich protein saliva is important to promote cell adhesion to tooth surface and
H NMR spectrum in the 6.0–7.6 ppm range.
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Fig. 4.
1
293
H NMR spectrum in the 1.0–5.5 ppm range.
the protein–polyphenols complexation in this model (Guggenheim et al., 2001; Spencer et al., 1988; Bennick, 2002). In order to compensate any loss in salivary proteins due to filtration, strains of Streptococcus salivarius was included in our biofilm model (Williams and Kraus, 1963). The Streptococcus salivarius is not an oral pathogen in mature dental biofilms, but it is ubiquitous in saliva and can produce large amounts of extracellular polymers when exposed to sucrose. This bacterium is a pioneer colonizing in dental plaque and creates favorable conditions so that other species can begin colonization (Marsh, 2005). Our biofilm model reflected an immature dental biofilm since we did not include Fusobacterium nucleatum, an oral bacterium related to bacteria co-aggregation and periodontal diseases. Nevertheless, an oral biofilm was created and the environmental conditions in our biofilm model were suitable for bacteria growth as shown by the data of the negative controls. Our results showed that the extract of Caesalpinia ferrea Martius has a good inhibitory effect upon oral microorganism in a planktonic state since MIC values ranged between 25 and 100 g/mL. A less pronounced effect was observed upon bacteria in a biofilm form since viable bacteria at 10−4 microbial dilution could be found. This last result was expected since bacterial biofilms are several hundred times more resistant than the MIC for the same bacteria in planktonic state. Our data suggest that the Amazonian tradition of using this plant for oral hygiene or infections is effective. Moreover, it can be speculated that the crude extract of this plant is active against free salivary oral bacteria in vivo. The biofilm model was a valuable attempt to simulate the plant mouthwash under in vivo conditions. Our findings support that the extract was effective but with lower activity when compared to a regular chlorhexidine mouthwash (1.2 mg/mL). However, for this experiment the extract was diluted in water in order to avoid interference in cell adhesion due to the high polarity of DMSO (Marshall et al., 1989). Thus, the plant extract in the biofilm assay was basically composed of water-soluble compounds and some biological activity could be compromised in this process. The 1 H NMR spectrum
confirms the presence of several chemical constituents including polyphenols and many other organic compounds (Cowan, 1999; Souza et al., 2006). The evident presence of tannins in the extract can certainly provoke salivary protein biding affecting salivary bacteria, but other active constituents in the extract are certainly contributing for this biological activity and other antimicrobial mechanisms of action. For instance, aiming to control dental caries, some alkaloids, flavonoids and tannins have recently proved to be effective against the same cariogenic bacteria species evaluated in our study (Badria and Zidan, 2004). Under clinical conditions the antibacterial effect of many natural products might be reduced due to constant salivary flow whereas chlorhexidine is less affected by saliva due to its high substantivity (Scheie and Petersen, 2004). Except for essential oils few natural compounds were tested in vivo for their ability to be present in the oral cavity for longer periods. Tea polyphenols molecules can be adsorbed and affect the acquired salivary pellicle (Joiner et al., 2004). This is an indication that polyphenols may interfere more on biofilm formation than on cells of a mature biofilm. In our study, a pre-exposure of the blocks to the plant extract was not tested and this effect could not be evaluated. This seems to be true since cocoa polyphenols reduces pH production without affecting Streptococcus mutans growth and that the crude extract of cranberry can affect the accumulation and polysaccharide composition of Streptococcus mutans biofilms (Duarte et al., 2006; Percival et al., 2006). The synergic or isolated activity of other constituents was not evaluated in our study, but this approach may render interesting results for controlling oral biofilm growth. For prophylactic purposes, many authors support that the active agents of antimicrobials should prevent biofilm formation without affecting the biological equilibrium within the oral cavity (Baehni and Takeuchi, 2003; Scheie and Petersen, 2004; Marsh and Percival, 2006). Since this is a characteristic of many organic compounds, medicine plants can certainly be a valuable source of oral antimicrobials (Cowan, 1999). This is particularly important when active
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compounds can be found in the fruits of plants such as Brazilian ironwood which are available in many local markets of the Amazonian region. Finally, our findings support that the fruits of Caesalpinia ferrea Martius have antibacterial activity against oral pathogens in planktonic or biofilm in vitro models. Acknowledgements This study was supported by CNPq 306234/2004-1 and FAPEAM—PROGRAMA POSVINC 1192-4-A. References Andrews, J.M., 2001. Determination of minimum inhibitory concentrations. Journal of Antimicrobial Chemotherapy 48 (Suppl. 1), 5–16. Bacchi, E.M., Sertie, J.A., Villa, N., Katz, H., 1995. Antiulcer action and toxicity of Styrax camporum and Caesalpinia ferrea. Planta Medica 61, 204–207. Badria, F.A., Zidan, O.A., 2004. Natural products for dental caries prevention. Journal of Medicinal Food 7, 381–384. Baehni, P.C., Takeuchi, Y., 2003. Anti-plaque agents in the prevention of biofilmassociated oral diseases. Oral Diseases 9 (Suppl. 1), 23–29. Bennick, A., 2002. Interaction of plant polyphenols with salivary proteins. Critical Reviews in Oral Biology and Medicine 13, 184–196. Borrás, M.R.L., 2003. Plantas da Amazônia: Medicinais ou mágica?—Plantas comercializadas no mercado Adolpho Lisboa. Editora Valer/Governo do Estado do Amazonas, Manaus. Carvalho, J.C.T., Teixeira, J.R.M., Souza, P.J.C., Bastos, J.K., dos Santos Filho, D., Sarti, S.J., 1996. Preliminary studies of analgesic and anti-inflammatory properties of Caesalpinia ferrea crude extract. Journal of Ethnopharmacology 53, 175–178. Cavalcante, R., 2008. As plantas medicinais na Odontologia: um guia prático. Expressão Gráfica, Rio Branco. Cowan, M.M., 1999. Plant products as antimicrobial agents. Clinical Microbiology Reviews 12, 564–582. Davies, D., 2003. Understanding biofilm resistance to antibacterial agents. Nature Reviews Drug Discovery 2, 114–122. Di Stasi, L.C., Hiruma-Lima, C.A., 2002. Plantas medicinais na Amazônia e na Mata Atlântica. Unesp, São Paulo. Duarte, S., Gregoire, S., Singh, A.P., Vorsa, N., Schaich, K., Bowen, W.H., Koo, H., 2006. Inhibitory effects of cranberry polyphenols on formation and acidogenicity of Streptococcus mutans biofilms. FEMS Microbiology Letters 257, 50–56. Graham, H.D., 1992. Stabilization of Prussian blue color in the determination of polyphenols. Journal of Agricultural and Food Chemistry 40, 801–805. Guggenheim, B., Giertsen, E., Schupbach, P., Shapiro, S., 2001. Validation of an in vitro biofilm model of supragingival plaque. Journal of Dental Research 80, 363–370. Guggenheim, B., Guggenheim, M., Gmur, R., Giertsen, E., Thurnheer, T., 2004. Application of the Zurich biofilm model to problems of cariology. Caries Research 38, 212–222. Huber, B., Eberl, L., Feucht, W., Polster, J., 2003. Influence of polyphenols on bacterial biofilm formation and quorum-sensing. Zeitschrift für Naturforschung C 58, 879–884. Joiner, A., Muller, D., Elofsson, U.M., Arnebrant, T., 2004. Ellipsometry analysis of the in vitro adsorption of tea polyphenols onto salivary pellicles. European Journal of Oral Sciences 112, 510–515. Kakiuchi, N., Hattori, M., Nishizawa, M., Yamagishi, T., Okuda, T., Namba, T., 1986. Studies on dental caries prevention by traditional medicines. VIII. Inhibitory effect of various tannins on glucan synthesis by glucosyltransferase from Streptococcus mutans. Chemical and Pharmaceutical Bulletin (Tokyo) 34, 720–725. Lorenzi, H., Matos, F.J.A., 2002. Plantas Medicinais no Brasil. Instituto Plantarum, Nova Odessa.
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