Reagents

All the chemical reagents that were used as solvents and standards in the procedures were of the analytical grade of the Sigma-Aldrich® brand (St. Louis, MO, USA); for the microbiological assays, the Mueller-Hinton Agar and Mueller-Hinton Broth culture media from Merck® (Darmstadt, Germany) were used.

Collection of samples

Four samples of native propolis were collected from beehives using a standard plastic mesh trap of the apiaries located in the municipalities of Torreón and Matamoros, Coahuila state, and the municipalities of Gómez Palacio and Lerdo, in the Durango state. These municipalities form part of the Comarca Lagunera, which is located in northern and central Mexico (25° 05' and 26° 54' N and 101º 40' and 104° 45' W). It has an average of 235 mm of rain, an elevation of 1,139 m and an average annual temperature of 18.6 °C (^{CIGEL, 2018}). All samples were kept in the darkness and stored at 4°C until use. With the samples from the four places, a mixture was formed, since that is how the population traditionally consumes them. This mixture was dried in a conventional oven (Linderberg/Blue, USA) at 55°C for 48 h. Moisture content was determined according the method 925.09 of the ^{AOAC (2014)}.

Obtaining extracts

The extracts of the propolis mixture were obtained by the classical method of maceration, by microwave-assisted extraction (^{Alonso-Castro et al., 2016}) and by ultrasound (^{Muñiz et al., 2013}).

Classical method of maceration. 20 g of the dry mixture of raw propolis was placed in a flask, and 200 mL of 70% ethanol was added; it was kept at 35°C at 150 rpm for 48 h on a shaker mixer (IKA KS 4000). Ultrasound-assisted extraction. 20 g of the dry mixture of crude propolis was placed in a flask, and 200 mL of ethanol (70%) was added; the flask was placed in an ultrasonic bath (Branson®) at 30°C to 300 W for 30 min. Microwave-assisted extraction. 20 g of the powdered dried stem of propolis was extracted with 315 mL of 70% ethanol using a closed system of microwave-assisted extraction (Multiwave 3000 Solv, Anton Paar, Graz, Austria). The extraction was performed at 70°C and 90 bars during 17 min. The extracts obtained by the three methods were filtered using filter paper 125 mm, No. 3 (Whatman®), and concentrated under reduced pressure in a rotary evaporator (BUGI®) to remove the solvent. The extracts were resuspended in distilled water and sterilized by membrane filtration with a 0.45 µm pore (Whatman®) and stored at 4°C until use. From the stock solution of each extract, the chemical groups were identified in each of them, and their antioxidant and antimicrobial activities were studied.

Phytochemical screening

The identification of chemical groups in the extracts of the propolis mixture was carried out by means of a phytochemical screening. Chemical groups corresponding to: alkaloids, sterols, unsaturations, sesquiterpenelactones, flavonoids, saponins, coumarins, and phenolic oxydrils were identified by colorimetric reactions according to the techniques described by ^{Ramman (2006)} and ^{Buvaneswari et al. (2011)}. Alkaloids: Mayer Test: To 0.5 mL of the extract, 6 drops of Mayer's reagent were added by carefully adding one side of the test tube. A white-creamy precipitate indicated the test as positive. Distilled water was used as a negative control and as positive control caffeine. Sterols: Liebermann-Burchard reaction: 1.0 mL of extract dissolved with 1.0 mL of chloroform was mixed with 1.0 mL of acetic anhydride and 0.5 mL of concentrated H₂SO₄. A series of color changes showed the presence of phytosterols. A blue-green ring indicated the presence of terpenoids. Distilled water was used as a negative control and as positive control almond oil. Unsaturations: A solution of 2% KMnO₄ was added dropwise in water to 0.5 mL of the extract; the test was positive if discoloration or formation of a brown precipitate was observed resulting from the formation of manganese dioxide. Distilled water was used as a negative control and as positive control olive oil. Sesquiterpenlactones: Baljet test: two solutions were used which were mixed in equal volumes before use. Solution A: 0.5 g of picric acid was placed in 50 mL of ethanol. Solution B: 5.0 g of NaOH in 50 mL of water was added. For the test, 50 μL of the extract and 4 drops of the reagent were added. The test was positive if the mixture becomes orange or dark red. Distilled water was used as a negative control and as positive control celery extract. Flavonoids: H₂SO₄ test: 50 μL of the extract and 10 μL of concentrated sulfuric acid were placed. The test was positive if yellow colorings were observed for flavones and flavonols; orange-icing for flavones; red-blue for chalcones and red-purple for quinones. Distilled water was used as a negative control and as positive control guava extract. Saponins: Shinoda test: Magnesium filings were added to 0.5 mL of the extract, and then a few drops of concentrated HCl were added. The test was considered positive with the appearance of colors orange, red, pink, and pink-blue to violet. Distilled water was used as a negative control and as positive control cassava. Coumarins: Sodium hydroxide test: 50.0 μL of the extract was placed and 10.0 μL of 10% NaOH solution was added; the test was positive when a yellow color appeared that disappeared when acidulating with 10.0 μL HCl. Distilled water was used with a negative control and as positive control cinnamon oil. Phenolic oxydriles: FeCl₃ test: 0.5 mL extract and 5% FeCl₃ drops were added together. The appearance of a red, violet-blue or green precipitate was considered positive. Distilled water was used with negative control and as positive control vanillin.

Antioxidant properties

Sample preparation. For the determination of the polyphenols and the antioxidant capacity of each propolis extract, 1 g was weighed and placed in a flask with 4.0 mL of water (ratio 1:5); the mixture was homogenized by constant stirring for 2 h at 500 rpm, protected against the light. It was then centrifuged at 10,000 rpm and stored in eppendorf tubes, and the respective analyses were performed immediately.

Determination of total content phenolic compounds

Total content phenolic compounds were determined using Folin-Ciocalteu reagent following the method described by ^{Singleton et al. (1999)}. From each propolis extracts, dilutions in ratio 1:10 were made with deionized water. 1 mL of this dilution was obtained and placed in a test tube, and 5 mL of Folin-Ciocalteu reagent (diluted 1:10 with deionized water) was added. It was allowed to stand for 8 min, and then 4 mL sodium bicarbonate solution (7.5%, w/v) was added, until a homogeneous mixture was obtained. The tubes were covered with foil to protect them from light and were incubated for 2 h at room temperature. The absorbance of each mixture was measured at 740 nm using a spectrophotometer (Genesys 10uv, USA).

To determine the total phenol content, the absorbance value obtained for each of the evaluated samples was replaced in the equation of a standard curve of gallic acid (^{Skerget et al., 2005}). The content of total phenolic compounds was expressed as milligrams of gallic acid equivalents per liter of extract (mg GAE•L^-1).

Estimation of total flavonoids content

Estimation of flavonoids was done as described by ^{Lagouri et al. (2014)}. The concentration of total flavonoids in the propolis extracts was measured spectrophotometrically at 415 nm. Half a milliliter of the sample was transferred to a 10-mL volumetric flask containing 2 mL of AlCl₃ (to 10% in ethanol) and 3 mL of CH₃COONa (50 g•L^-1 in ethanol). The controls contained all the reagents except for the extract. After 2.5 h at 20°C, the absorbance was measured at 440 nm. The same procedure was repeated for the standard quercetin solutions (25-250 ppm), and the results were expressed in milligrams of quercetin per gram of propolis and were presented as means of triplicates.

Antioxidant activity

Was performed according to the methodology described by ^{Brand et al. (1995)}, which it is based on inhibition of the stable radical 2,2-Diphenyl-1-picrylhydrazyl (DPPH). A calibration curve was done from a methanolic stock solution of DPPH radical, with a concentration of 20 mg•L^-1 and the (±)-6-Hydroxy-2,5,7,8-tetramethylchromane-2-carboxylic acid (Trolox reagent). In a Falcon tube, 990 μL of the DPPH radical and 10 μL of the solution of the ethanolic extract of propolis added, the mixture was shaken with vortex and allowed to stand for 30 min in the absence of light, and its absorbance was read in a spectrophotometer at 517 nm.

Antibacterial activity

The antibacterial activity was studied from the inhibitory effect of each extract against Staphylococcus aureus ATCC 35556, Escherichia coli ATCC 25922, Salmonella typhi ATCC 14028 and Proteus mirabilis ATCC 9150. From the revitalized strains, the inoculum was prepared as described by ^{Cockerill et al. (2012)}. The percentage of activity of each extract and the rate of bacterial susceptibility were also calculated.

Inhibitory effect

The inhibitory effect of each extract was measured from the zones of inhibition observed when using the disk diffusion technique (^{Souto et al., 2006}). 10 μL (1x10⁶ CFU) of bacteria were inoculated on the surface of Muller-Hinton agar. After that, filter paper discs (Whatman®) of 6.0 mm diameter were impregnated with 10 μL of each extract. Moxifloxacin (50 mg), a broad-spectrum antibiotic, was used as a positive control, and destilled water was used as a negative control. All tests were performed in triplicate.

Percentage of activity

The percentage of activity (equation 1) indicates the total antimicrobial potency of each extract (^{López-Malo et al., 2005}).

Index bacterial susceptibility (IBS)

The IBS (Equation 2) shows the number of microorganisms susceptible to the extract, assessing ranges from 0 (resistance extract all samples) to 100 (susceptible to the whole extract) (^{López-Malo et al., 2005}; ^{Panghal et al., 2011}).

Statistical analysis

Data were statistically analyzed by one-way analysis of variance (ANOVA) to test differences between extraction methods, and a (Student-Newmal-Keuls) comparison of means was used to determine the differences between groups when necessary. Statistical analyses were conducted using Statistica software. Differences were considered statistically significant at 95% of confidence level.

Phytochemical screening

The moisture of the dry propolis mixture from which the extracts was 1.66% ± 0.29. The phytochemical screening of the hydroalcoholic extract of propolis revealed the presence of important secondary metabolites. Likewise, there was no difference in the presence of metabolites analyzed in relation to the extraction method used.

The bioclimate and flora of the Comarca Lagunera, whose vegetation is composed of Prosopis spp., Abies alba Mill., Acacia farnesiana (L.) Willd, Phoenix spp. and Larrea tridentata (Sessé and Moc. ex DC.) Coville, varieties (^{Estrada-Rodríguez, 2004}), favor the presence of bioactive compounds in propolis. The result of phytochemical screening of propolis samples coincides with that reported by ^{Soltani et al. (2017)}, who by means of gas chromatography and mass spectrometry, detected: aromatic acid ester, saturated hydrocarbon alkaloids, saturated hydrocarbons, and aromatic heterocyclic organic compounds, in ethanolic extracts of Algerian propolis. Among the numerous groups of substances identified in propolis samples from different localities, the most common are aromatic acids and esters, chalcones, flavonoids, terpenoids, and waxy acids. Furthermore, most of the biological activities of propolis have been attributed to these compounds, especially the flavonoids (^{Trusheva et al., 2006}). A significant number of reports describe its anti-inflammatory, immunomodulatory, antioxidant, anticancer, and hepatoprotective properties, as well as many other biological activities of propolis (^{Borges et al., 2011}).

Total content phenolic compounds and antioxidant activity

When quantifying the content of phenols in the extracts of a mixture of propolis obtained by maceration, assisted by ultrasound and by microwaves methods, it was observed that the method used to obtain the extracts does affect the content of phenols and flavonoids as well as the antioxidant capacity on which phenols have more influence than flavonoids (Tabla 1). The traditional maceration method was better for the extraction of phenolic compounds because it was observed a 40 and 28% more with respect to the other 2 methods; however, the extraction assisted by ultrasound was better to extract falvonoids, since it was obtained 54 and 52% more than the methods of maceration and assisted by microwaves respectively. Although the antioxidant capacity of the extract obtained by ultrasound was reduced by 8%, in relation to the other two methods, which suggests that the antioxidant capacity it is not only due to flavonoids, but to a synergistic effect with other bioactive compounds present in the sample

The interest in quantifying phenolic compounds and flavonoids present in propolis is because they are responsible for the physiological activity of this product (^{Delgado et al., 2015}). ^{Farré et al. (2004)} consider that the phenolic compounds act synergistically with flavonoids, and of these, quercetin is one of the flavonoids of interest to quantify.

Antibacterial activity

The extracts obtained by the three methods showed 100% of antimicrobial activity and IBS was of 100%. The inhibitory effect of the extracts (Table 2) varied, not only regarding the extraction method, but also depending on the bacteria studied. It was observed that the inhibitory effect was less for E. coli than for the other studied bacteria.

There was no significant difference (p <0.5) between the diameters of the zone of inhibition of extracts obtained by different techniques, which supports the way in which the extract is traditionally used, that is, by maceration. The least sensitive microorganism was S. typhi, and the most sensitive was E. coli. Although the three extracts were active against the bacteria studied, none of them was as effective as the antibiotic used as a positive control. These results are similar from those reported by ^{Ertürk et al. (2016)} and for ^{Koo et al. (2000)}, who reported that the ethanolic extract of crude extract of propolis was found the most sensitive microorganism to propolis was E. coli in the Gram negative group. This activity can be a synergy between flavonoids, apigenin, chrysin, or other components in raw propolis samples. According to ^{Hegazi et al. (2014)}, the propolis samples show a different antimicrobial activity and attribute it to the complex composition of the resin. In general, propolis is used to treat various diseases in humans. One of the most important properties of propolis is its antimicrobial activity, which is attributed mainly to flavonoids, the main biologically active compound in propolis, and it is a substance that has potential for the treatment of conditions caused by different microorganisms (^{Tolosa and Cañizares, 2002}). Several trials have demonstrated the antibacterial activity of ethanolic extracts against Staphylococcus aureus, E. coli and Pseudomonas spp. (^{Ortega et al., 2011}; ^{Díaz-Mena et al., 2000}), and it is currently investigated whether the propolis can be used as a natural food preservative (^{Vargas-Sánchez et al., 2013}). ^{Dimov et al. (1992)} correlated the antibacterial activity of propolis with the presence of flavonoids; and ^{Farnesi et al. (2009)} reported that the bactericidal effect of these compounds on the metabolic disturbance of ion channels due to paired phosphorylation / dephosphorylation reactions. The antimicrobial activity is also attributed to phenolic acids: ferulic, caffeic and cumaric, bioactive compounds found in propolis samples by ^{Popova et al. (2010)}. The presence of phenolic compounds in the samples of propolis explain their antimicrobial activity, which is supported by ^{Marcucci (1995)}, who reported that caffeic acid, as well as benzoic acid and cinnamic acid act on the microbial membrane, causing structural damage to the cells. Until now, no single propolis component has shown to possess more antibacterial properties than that of the total extract. Some authors cite the highly complex and variable composition of propolis as a reason for its antimicrobial activity and the data gathered to date suggest that such activity can be linked to multiple targets, with several constituents acting in synergy (^{Scazzocchio, 2006}). Propolis affects the cytoplasmic membrane, inhibits bacterial motility and enzyme activity, exhibits bacteriostatic activity against different bacterial genera and can act as a bactericide at high concentrations (^{Mirzoeva, 1997}).