The aim of this study was to investigate the antimicrobial activity of ethanolic extract and fractions
of Brazilian green propolis (BGP) collected by bees from Baccharis dracunculifolia against 16 oral
pathogenic microorganisms. BGP was examinated by Reversed-Phase High-Performance Liquid
Chromatography (RPHPLC) and its absorption spectra was assessed using UV-Spectrophotometer.
Identification of flavonoids and other chemical constituents were carried out using authentic
standards. Antimicrobial activity was evaluated by agar diffusion and dilution method. The results
indicate that all microorganisms tested were susceptible to BGP. None of the essayed fractions
(Coumaric acid, Kaempferol, Pinobanskin-3-acetate, Chrysin, Galangin, Kaempferide, and Artepillin
C) was more active than the extract, suggesting a synergistic effect of propolis constituents for the
antimicrobial activity.
Propolis is a natural resinous hive product used by the bees to protect the hive against the invasion of
microorganisms and is considered to be a natural antibiotic1. It has also been used extensively in folk
medicine by the Brazilian population for several years. Many beekeepers, pharmacists and local
laboratories in Brazil produce a great variety of propolis derivatives for medicinal use. Beekeepers
commonly chew raw propolis to treat mouth and upper digestive track infections. Available literature
indicates that few antimicrobial studies have been carried out using Brazilian propolis and there are
only a few reports documenting the chemical constituents and their biological activities2,3. The aim of
this study was to evaluate the antibacterial and antifungal activity of Brazilian green propolis extract
and fractions against Candida spp., Gram positive and Gram negative oral pathogenic bacteria.
Crude Brazilian green propolis was obtained from an apicultural region of Minas Gerais State, Brazil
by PharmaNectar®(REF. LOT. SBN97). 100g of crude propolis was kept in a freezer for 24 h and
powdered in a blender. After dissolving, ethanolic extracts of Brazilian green propolis (BGP) was
filtered through Sigma nº1 filter paper. The filtered extract was concentrated under vacuum to furnish
62g of a crude extract. Analysis of flavonoids from ethanolic extracts of bud and unexpanded leaf
exudates and ethanolic extracts of propolis was performed by RPHPLC2 with a chromatograph
equipped with a YMC PACK ODS-A column (RP-18, column size 4.6 x 250 mm; particle size 5μm)
and Photodiode Array Detector (SPD- M10A, Shimazu Co., Japan). The column was eluted by using
a linear gradient of water (solvent A) and methanol (solvent B) starting with 30% B (0-15 min), and
increasing to 90% B (15-75min), held at 90% B (75-95 min) and decreasing to 30% B (95-105 min)
with a solvent flow rate of 1 mL/min and detection with a diode array detector. Chromatograms were
recorded at 268nm. The ethanolic extract of propolis was measured the absorption spectra, using UVSpectrophotometer.
Identification of flavonoids and other chemical constituents were carried out using authentic
standards purchased from Extrasythese Co. (France). The authentic standard of 3,5-dipremyl-4-
hydroxycinnamic acid (artepillin C) was donated from Hayashibara Biochemical Laboratory
(Okayama, Japan). The authentic standard of Pinobanksin and Pinobanksin-3-acetate were donated
from Dr. E. Wollenweber (Institue für Botanik, Technishe Hochschule Darmstadt, Germany). The
minimal inhibitory concentration (MIC) was defined as the lowest concentration of propolis in which
no bacterial growth was detected. Determination of MICs by the agar dilution method was
performed, following the serial concentrations of BGP and different fractions were achieved (%v/v)
in plates containing Brucella agar (Oxoid), as follows: 0.1, 0.2, 0.4, 0.8, 1.75, 3.5, 7.0 and 14.0. Each
antimicrobial test also included plates containing the culture medium plus ethanol, in order to obtain
a control of the solvent antibacterial effect4,5. The antimicrobial and antifungal susceptibility test for
Streptococcus mutans (ATCC 70069), Strepetococcus sanguis (ATCC 10557), Lactobacillus casei
(ATCC 393), Tanerella forsythensis (ATCC 700191), Bacteroides fragilis (ATCC 25285),
Staphylococcus aureus (ATCC 12692), Fusobacterium necrophorum (ATCC 25286), Actinobacillus
actinomycetemcomitans (ATCC 33384), Porphyromonas gingivalis (ATCC 33277), Fusobacterium
nucleatum (ATCC 23726), C. albicans (ATCC 18804), C. tropicalis (ATCC 750), C. glabrata
(ATCC 2001), C. parapsilosis (ATCC 22019), C. krusei (ATCC 2340) and C. guilliermondii (ATCC
201935) were studied with reference microdilution method following the NCCLS M27-AZ Standards
by using RPMI 1640 medium (Sigma-USA) with L-glutamine and phenol red and without sodium
bicarbonate6, 7. Yeast suspension were inoculated into microplate wells which contained 1/64-1/8000
dilution of BGP and fractions solutions. Microplates were evaluated after incubation at 37ºC for 48 h.
Sterile blank disks (CECON - São Paulo - Brazil) were soaked in 20 μl of the BGP solution, 20μl of
each component Coumaric acid, kaempferol, Pinobanskin-3-acetate, chrysin, galangin, kaempferide,
and artepillin C, and applied to the agar surface previously seeded with the microorganism.
Positive and negative controls of the discs containing 30μg of tetracycline, Nystatin 30mg, and 20μl
of Ethanol 93,2ºC were used. After 48 hours of incubation at 37ºC, the diameters of the inhibition
zones were measured and compared. The results of the diameters of the inhibition zones were
reported as Means + Standard Deviation (M±SD). The inhibitory ability of the various propolis
solutions tested on the oral pathogenic bacteria and fungi was compared with nonparametric Kruskal-
Wallis test. Differences of the level p<0.05 were considered to be significant.
