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An efficient chemical analysis of phenolic acids and flavonoids in raw propolis by microwave-assisted extraction combined with high-performance liquid chromatography using the fused-core technology
Journal of Pharmaceutical and Biomedical Analysis 81–82 (2013) 126–132
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An efficient chemical analysis of phenolic acids and flavonoids in raw propolis by microwave-assisted extraction combined with high-performance liquid chromatography using the fused-core technology Federica Pellati ∗ , Francesco Pio Prencipe, Davide Bertelli, Stefania Benvenuti Department of Life Sciences, University of Modena and Reggio Emilia, Via G. Campi 183, 41125 Modena, Italy
a r t i c l e
i n f o
Article history: Received 8 February 2013 Received in revised form 3 April 2013 Accepted 5 April 2013 Available online xxx Keywords: Propolis Flavonoids Phenolic acids Microwave-assisted extraction Fused-core HPLC
a b s t r a c t A closed-vessel microwave-assisted extraction (MAE) technique was optimized for the first time for the extraction of polyphenols from raw propolis. The results obtained by means of response surface experimental design methodology showed that the best global response was reached when the extraction temperature was set at 106 ◦ C, the solvent composition close to EtOH–H2 O 80:20 (v/v), with an extraction time of 15 min. In comparison with other techniques, such as maceration, heat reflux extraction (HRE) and ultrasound-assisted extraction (UAE), the extraction with MAE was improved by shorter extraction time and lower volume of solvent needed. The HPLC analyses of propolis extracts were carried out on a fused-core Ascentis Express C18 column (150 mm × 3.0 mm I.D., 2.7 m), with a gradient mobile phase composed by 0.1% formic acid in water and acetonitrile. Detection was performed by DAD and MS. The method validation indicated that the correlation coefficients were >0.999; the limit of detection was in the range 0.5–0.8 g/ml for phenolic acids and 1.2–3.0 g/ml for flavonoids; the recovery range was 95.3–98.1% for phenolic acids and 94.1–101.3% for flavonoids; the intra- and inter-day %RSD values for retention times and peak areas were ≤0.3 and 2.2%, respectively. The quali- and quantitative analysis of polyphenols in Italian samples of raw propolis was performed with the validated method. Total phenolic acids ranged from 5.0 to 120.8 mg/g and total flavonoids from 2.5 to 168.0 mg/g. The proposed MAE procedure and HPLC method can be considered reliable and useful tools for the comprehensive multi-component analysis of polyphenols in propolis extracts to be used in apitherapy. © 2013 Elsevier B.V. All rights reserved.
1. Introduction Propolis is a resinous material collected by honeybees (Apis mellifera L.) from several tree species [1]. In regions with temperate climate, the resin is collected mainly from the buds and cracks in the bark of Poplar trees [2]. Propolis is characterized by a series of biological properties, such as antibacterial, antiviral, antifungal, anti-inflammatory, antioxidant, antiproliferative, immunostimulating [1]. Typical applications of propolis include herbal products for cold syndrome and dermatological preparations [3]. Propolis extracts are also used to prevent and treat oral inflammations [3].
Abbreviations: MAE, microwave-assisted extraction; HRE, heat reflux extraction; UAE, ultrasound-assisted extraction; CCD, central composite design; DMCA, 3,4dimethyl caffeic acid; CAPE, caffeic acid phenylethyl ester. ∗ Corresponding author. Tel.: +39 059 205 5144; fax: +39 059 205 5131. E-mail address: [email protected] (F. Pellati). 0731-7085/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.jpba.2013.04.003
The chemical composition of propolis is known to be very complex and influenced by the geographic origin [3]. In propolis from temperate zones, the most representative biologically active compounds are polyphenols, including phenolic acids and flavonoids [2,3]. Raw propolis cannot be used as crude material, but it must be purified by solvent extraction to remove the useless material and preserve the active polyphenolic fraction [2,3]. Hydroalcoholic solvents have been described as the most suitable media for the extraction of biologically active phenolic components from raw propolis [2]. Regarding the extraction techniques used for the chemical analysis of raw propolis, maceration at room temperature and heat reflux extraction (HRE) have been widely applied [2,4]. However, conventional extraction methods are often timeconsuming and require high solvent volumes, raising process costs and reducing environmental sustainability [5,6]. For this reason, the exploration of innovative extraction techniques that could solve some of the above mentioned problems has been considered [6]. In this ambit, microwave-assisted extraction (MAE) represents a
F. Pellati et al. / Journal of Pharmaceutical and Biomedical Analysis 81–82 (2013) 126–132
reliable alternative to traditional extraction techniques [7] and it has been widely applied to the extraction of flavonoids from plant material [8]. Several examples in the ambit of phytochemical analysis suggested that MAE has some considerable merits, such as shorter extraction time, higher extraction yield and less solvent consumption compared to conventional extraction methods [7]. Different types of MAE systems can be applied to the extraction of phenolics from natural products, including microwave ovens operating at the atmospheric pressure and closed-vessel equipments under controlled temperature and pressure [7,8]. In a previous study, Trusheva et al. [9] used a multimodal household microwave oven at 800 W for the extraction of polyphenols from propolis; the authors have observed the presence of unwanted compounds in the extracts and a decrease in the amount of polyphenols, probably due to the very high microwave power applied [10]. In addition, the use of domestic ovens without technical modifications should be avoided, because of the low safety, reliability and reproducibility which characterize these equipments [7]. In this study, a response surface experimental design methodology coupled with central composite design (CCD) was applied for the first time to optimize the extraction of polyphenols from raw propolis using MAE. To the best of our knowledge, closed-vessel MAE with a system operating at controlled temperature and low irradiation power has never been applied before to the extraction of phenolic compounds from raw propolis. The extraction efficiency of MAE was compared with that of other techniques previously described in the literature [2,4], including maceration, HRE and ultrasound-assisted extraction (UAE), in terms of yield, time and solvent usage. The analysis of polyphenols in propolis extracts was carried out by an improved RP-HPLC method using a fused-core C18 column. The method was fully validated and its practical applicability was demonstrated by the analysis of Italian samples of raw propolis to provide a complete chemical analysis of their biologically active constituents.
2. Materials and methods 2.1. Propolis samples, chemicals and solvents Nine samples of raw propolis (indicated in the text as RP-1/RP-9) were collected from Apis mellifera hives located in different Italian regions in Spring 2012 and stored at −20 ◦ C until chemical analysis. The frozen samples were finely powdered using a mortar and a pestle before the extraction procedure. Sample labeled as RP-1 was used for the optimization of the extraction conditions, because it was available in higher amount. Caffeic acid, p-coumaric acid, ferulic acid, quercetin, pinocembrin, cinnamic acid, apigenin, kaempferol and galangin were from Sigma–Aldrich–Fluka (Milan, Italy). Isorhamnetin and luteolin were from Roth (Karlsruhe, Germany). Chrysin was from Extrasynthese (Genay, France). The purity of standard compounds was always higher than 97%, as confirmed by HPLC-DAD analysis. HPLC-grade acetonitrile (ACN), formic acid and analytical grade absolute ethanol (EtOH) were from Sigma (Milan, Italy). Water was purified using a Milli-Q Plus185 system from Millipore (Milford, MA, USA).
2.2. HPLC-DAD conditions HPLC analyses were performed on an Agilent Technologies (Waldbronn, Germany) modular model 1100 system consisting of a vacuum degasser, a quaternary pump, an autosampler, a thermostatted column compartment and a diode array detector (DAD).
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The chromatograms were recorded using an Agilent Chemstation for LC and LC–MS systems (Rev. B.01.03). An Ascentis Express C18 column (150 mm × 3.0 mm I.D., 2.7 m, Supelco, Bellefonte, PA, USA) was used for the HPLC analysis. The mobile phase was composed by 0.1% formic acid in H2 O (A) and ACN (B). The gradient elution was modified as follows: 0–3 min 20% B, 3–10 min from 20 to 30% B, 10–40 min from 30 to 40% B, 40–50 min from 40 to 60% B, 50–60 min from 60 to 80% B, 60–65 min from 80 to 50% B. The post-running time was 10 min. The flow rate was 0.6 ml/min. The column temperature was set at 30 ◦ C. The sample injection volume was 2 l. The DAD acquisitions were carried out in the range 190–450 nm and chromatograms were integrated at 265 nm (for chrysin and galangin), 290 nm (for cinnamic acid and pinocembrin), 320 nm (for caffeic acid, p-coumaric acid and ferulic acid), 338 nm (for apigenin and luteolin) and 370 nm (for quercetin, isorhamnetin and kaempferol). Three injections were performed for each sample. 2.3. HPLC–ESI–MS and MS2 conditions HPLC-ESI-MS and MS2 analyses were carried out using an Agilent Technologies modular 1200 system, equipped with a vacuum degasser, a binary pump, an autosampler, a thermostated column compartment and a 6310A ion trap mass analyzer with an ESI ion source. The HPLC column and the applied chromatographic conditions were the same as reported for the HPLC-DAD system. The flow rate was split 3:1 before the ESI source. The experimental parameters were set as follows: the capillary voltage was 3.5 kV, the nebulizer (N2 ) pressure was 32 psi, the drying gas temperature was 350 ◦ C, the drying gas flow was 11 l/min and the skimmer voltage was 40 V. Data were acquired by Agilent 6300 Series Ion Trap LC/MS system software (version 6.2). The mass spectrometer was operated in the full-scan positive and negative ion modes in the m/z range 100–1000. MS2 spectra were automatically performed with helium as the collision gas by using the SmartFrag function. 2.4. Methods for the extraction of phenolics from raw propolis Four methods were applied and compared in order to obtain a high extraction efficiency of phenolic components from raw propolis. In each case, the applied sample-to-solvent ratio was 1:10 (w/v), as previously described in the literature [2,9]. The extraction procedure was repeated twice for each method. 2.4.1. Maceration, heat reflux extraction (HRE) and ultrasound-assisted extraction (UAE) Maceration extraction was performed on a weighed amount of sample (1.00 g) with 10 ml of EtOH–H2 O (80:20, v/v) at room temperature for 24 h under stirring. The HRE was carried out by refluxing a weighed amount of sample (1.00 g) with 10 ml of EtOH–H2 O (80:20, v/v) at 70 ◦ C for 1 h under stirring using a water bath. The UAE was performed on a weighed amount of sample (1.00 g) with 10 ml of EtOH–H2 O (80:20, v/v) at 70 ◦ C for 1 h using an ultrasonic bath (Sonorex RK-100H, Bandelin, Berlin, Germany). At the end of each of these extraction procedures, the mixture was centrifuged at 4000 rpm for 5 min. The supernatant solution was then paper filtered in a vacuum into a 10 ml volumetric flask and the solvent was added to the final volume. One ml aliquot of each propolis hydroalcoholic extract was diluted 1:5 (v/v) with the extraction solvent in a volumetric flask, filtered through a 0.45 m PTFE filter into a HPLC vial and injected into the HPLC system. Three injections were performed for each sample.
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Table 1 Factor levels and experimental domain applied to optimize the MAE experimental conditions. Factor
Solvent composition (EtOH–H2 O ratio, v/v) Temperature (T, ◦ C) Time (t, min)
Solvent composition (EtOH–H2 O ratio, v/v) Temperature (T, ◦ C) a b
First experimental domain −˛a
−1
0
1
˛a
49.7 66.4 6.6
60.0 80.0 10.0
75.0 100.0 15.0
90.0 120.0 20.0
100.2 133.6 23.4
Second experimental domain −˛b −1
0
1
˛b
53.8 71.7
75.0 100.0
90.0 120.0
96.2 128.3
60.0 80.0
˛ = 1.682. ˛ = 1.414.
2.4.2. Microwave-assisted extraction (MAE) Regarding MAE, a weighed amount of ground sample (0.50 g) was extracted with 5 ml of solvent in a 10 ml glass vessel using a monomode focused microwave apparatus with a closed-vessel system (Discover instrument, CEM, Metthews, NC, USA). The microwave equipment used in this study allowed extraction temperature, pressure and time to be programmed. Microwave power, with a maximum of 300 W and a magnetron frequency of 2450 MHz, and pressure were dynamically adjusted by temperature and power feedback control. The instrument was able to provide continuous non-pulse microwave heating, focusing on a single cavity where the sample was placed. During the extraction, magnetic stirring was applied to homogenize the sample. The optimized experimental conditions were set as follows: extraction solvent EtOH–H2 O (80:20, v/v), extraction temperature at 106 ◦ C and extraction time of 15 min. After the extraction time had elapsed, the vessel was allowed to cool at room temperature before opening. After centrifugation for 5 min at 4000 rpm, the supernatant solution was paper filtered in a vacuum into a 5 ml volumetric flask and the solvent was added to the final volume. Further processing of the samples prior to HPLC analysis was as described above.
2.5. Experimental design In order to optimize the extraction parameters of phenolics from raw propolis using MAE, an experimental design was applied. The optimization of MAE conditions was performed using a CCD (with ˛ = 1.682), which was based on a 23 factorial design (each experiment performed in triplicate) plus six axial points (each experiment performed in triplicate) plus six replicates in the center of the design. For MAE optimization, the variables chosen were the solvent composition (EtOH–H2 O ratio, v/v), the extraction temperature (T, ◦ C) and the extraction time (t, min). The factor levels and experimental domain are shown in Table 1. In particular, the solvent composition varied from EtOH–H2 O 50:50 (v/v) to 100% EtOH, covering the concentration range characterized by a high extraction yield for polyphenols from propolis [11]. The extraction temperature varied from 66 to 134 ◦ C, avoiding higher temperature that could cause the degradation of flavonoids, as described by Liazid et al. [12]. Finally, the extraction time varied from 7 to 23 min in order to obtain a fast extraction method. A total of forty-eight experiments were performed in randomized order. In view of the results of these experiments, the two most significant variables (EtOH–H2 O volume ratio and temperature) were selected to perform a second CCD with ˛ = 1.414, based on a 22 factorial design (each experiment performed in duplicate) plus four axial points (each experiment performed in duplicate) plus four replicates in the center of the design. The experimental domain was defined taking into account the results obtained from the first
CCD with some adjustments (Table 1), i.e. the solvent composition varied from EtOH–H2 O 54:46 (v/v) to 96% EtOH and the extraction temperature from 72 to 128 ◦ C. The extraction time was kept constant at 15 min. Twenty experiments were performed in randomized order. The analysis of the second CCD was performed by response surface methodology; this surface is approximated by a polynomial function, using a quadratic model, and represents a good description of the relationship between the experimental variables and the response within a limited experimental domain. Statistical analysis was performed using Statistica v. 6.1 (Statsoft Inc., Tulsa, OK, USA).
2.6. HPLC-DAD method validation The validation of the HPLC-DAD method was performed to show compliance with international requirements for analytical techniques for the quality control of pharmaceuticals (ICH guidelines) [13]. Regarding linearity, the stock solution of each compound commercially available (caffeic acid, p-coumaric acid, ferulic acid, quercetin, cinnamic acid, apigenin, kaempferol, isorhamnetin, luteolin, chrysin, pinocembrin and galangin) was prepared as follows: an accurately weighed amount of each pure standard compound (2–6 mg) was placed into a 10 ml volumetric flask; EtOH was added and the solution was diluted to volume with the same solvent. The external standard calibration curve was generated using seven data points. Injections were performed in triplicate for each concentration level. The calibration curve was obtained by plotting the peak area of the compound at each level versus the concentration of the sample. For each compound, the limit of detection (LOD) and the limit of quantification (LOQ) were experimentally verified by HPLC analysis of serial dilutions of a standard solution to reach a signal-to-noise (S/N) ratio of 3 and 10, respectively. The accuracy of the analytical procedure was evaluated using the recovery test. This involved the addition of a known quantity of standard compound to half the sample weight of raw propolis (sample RP-1) to reach 100% of the test concentration. The fortified samples were then extracted using the optimized MAE conditions and analyzed by the proposed HPLC method. The precision of the extraction technique was validated by repeating the extraction procedure on the same sample of raw propolis (RP-1) at the best MAE conditions. An aliquot of each extract was then injected and quantified. The extraction was performed in duplicate on three different days with newly prepared mobile phase and samples. The precision of the chromatographic system was tested by performing intra- and inter-day multiple injections of a hydroalcoholic extract of propolis (RP-1) and then checking the %RSD of retention times and peak areas. Six injections were performed each day for three consecutive days.
F. Pellati et al. / Journal of Pharmaceutical and Biomedical Analysis 81–82 (2013) 126–132 mAU 700
129
1 21
600 24
5
500
7+8
400
23
300
26
11
2 4
31
200
36
20
16
22
25
27
3
34+35 9
100 6
10 1314 12 15
17
32
28
18
37
33
19 29
38
30
0 0
10
20
30
40
50
min
Fig. 1. Chromatogram obtained by RP–HPLC–DAD analysis of a representative propolis extract (sample RP-6) at 290 nm on the fused-core column. Experimental conditions as in Section 2.2. For peak identification, see Table 2.
The stability of the analytes during MAE was evaluated with reference standard solutions subjected to HPLC analysis before and after the extraction treatment under the optimized conditions. Stability was also tested using propolis extracts stored in amber glass flasks at 4 ◦ C and at room temperature (about 25 ◦ C) and analyzed every 12 h. 3. Results and discussion 3.1. HPLC method development and optimization An initial comparison of fully porous and fused-core C18 columns for the HPLC analysis of polyphenols in raw propolis was carried out. In particular, a method initially developed on a fully porous Ascentis C18 column for the analysis of commercially available propolis extracts [4] was transferred to a fused-core Ascentis Express C18 column. A parallel careful adaptation of the experimental conditions was performed. In this way, the total analysis time was decreased from 92 to 65 min, with a consequent advantage in terms of time and solvent usage. Under these conditions, the highest observed backpressure was about 300 bar, within the pressure limit of conventional HPLC systems. The HPLC-DAD analysis at 290 nm of a hydroalcoholic extract obtained from a representative sample of raw propolis (RP-6) indicated a very complex composition, as shown in Fig. 1. A total of 38 compounds were identified in propolis samples, on the basis of their UV, MS and MS2 data, which were compared with those of the reference standards and with the literature [4]. Taking into account the complexity of the sample, the chromatographic separation can be considered satisfactory. 3.2. Optimization of extraction conditions Four methods were evaluated in order to obtain an efficient and rapid extraction of polyphenols from raw propolis. The comparison was carried out between conventional extraction methods (including maceration, HRE and UAE) [2,4] and MAE, which represents a more innovative extraction procedure. Since closed-vessel MAE has never been investigated for the extraction of phenolics from raw propolis, its experimental parameters were optimized. For a closed-vessel MAE system, the variables that could affect the extraction efficiency include temperature, solvent composition, time, microwave power, pressure and sample-to-solvent ratio [8,14,15]. In this study, the attention was focused on temperature, solvent composition and time, which are
recognized as representative parameters affecting the efficiency of MAE [8]. It should be noted that the microwave power was not chosen for the optimization of the extraction conditions, because the closed-vessel MAE system used in this work regulates the microwave power depending on the selected temperature. Furthermore, in view of the solvent mixtures used in this study, the microwave system reached the desired temperature at low irradiation power: in fact, under the applied conditions, the microwave power never exceeded 120 W, thus avoiding a possible degradation of flavonoids due to high irradiation energy [10]. Pressure is another important experimental variable for closed-vessel MAE, but it is directly dependent on temperature [15] and, therefore, it was not chosen for method optimization. Other parameters, such as sample amount and solvent volume were kept constant, with a 1:10 (w/v) ratio [2,9]. Response surface experimental design methodology coupled with CCD is a useful tool to optimize procedures depending on several variables [16] and, therefore, it was applied in this study. A response based on the total amount (mg/g) of the most representative propolis constituents under the conditions of the design was studied. The experimental values of the factors chosen for the optimization of MAE parameters were selected in order to cover a wide range of conditions. The data obtained were evaluated by ANOVA, the level of significance being set at 5%, to evaluate the statistical significance of each factor and interactions between the different factors. The quadratic term of the solvent composition was the most significant parameter (at p < 0.05) on the extraction yield of the compounds of interest. The linear and the quadratic term of the extraction time, as well as its interaction with the other factors, did not show a significant influence on the efficiency of the extraction process. A second CCD was thus set up, focusing on the solvent composition and the extraction temperature. As a confirmation of what observed with the previous CCD, the linear and the quadratic term of the solvent composition were the most significant parameters (at p < 0.05), having a strong influence on the extraction yield of phenolics from raw propolis. The interaction between solvent composition and extraction temperature was also found to be significant in the extraction procedure. Fig. 2 shows the response surface plot for the total amount of the extracted compounds versus solvent composition and extraction temperature, with an extraction time equal to 15 min. The experimental results showed that the best global response, within the range studied, was reached when the extraction temperature was set at 106 ◦ C, the extraction solvent close to EtOH–H2 O
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Concentration ( µg/mL)
400
300
Before MAE
200
After MAE 100
Galangin
Pinocembrin
Chrysin
Isorhamnetin
Kaempferol
Apigenin
Cinnamic acid
Quercetin
Ferulic acid
p-Coumaric acid
Caffeic acid
0
Fig. 4. Stability study of phenolic acids and flavonoids used as propolis standards subjected to optimized MAE conditions.
by HPLC analysis. The results indicated that the amount of polyphenols obtained from the residue after the first extraction was below the LOQ values in all cases. These results indicated that MAE represents an attractive sample preparation method and has good potential for the extraction of polyphenols from raw propolis. Fig. 2. Response surface plot for total amount (mg/g) of six representative polyphenolic compounds extracted from raw propolis versus solvent composition (EtOH–H2 O ratio, v/v) and extraction temperature (T, ◦ C), with a constant extraction time (t) equal to 15 min.
80:20 (v/v), with an extraction time of 15 min. These values were therefore selected for the extraction of polyphenols from raw propolis in all the subsequent analyses. The total amount of representative propolis phenolic compounds extracted under optimized MAE conditions was then compared with those obtained with maceration, HRE and UAE. As shown in Fig. 3, the MAE technique allowed to obtain the yield of other methods in only 15 min. With the assistance of the fast heating of microwave irradiation, the extraction with MAE was improved by a reduced extraction time and half volume of solvent needed. The composition of propolis extracts processed by closed-vessel MAE was monitored in detail by HPLC-DAD at five different wavelengths, by MS and MS2 , and the results indicated the absence of unwanted compounds. In addition, degradation products were not observed in the HPLC chromatograms. An investigation on the residue composition after MAE was also performed, by carrying out further extraction steps with the same solvents and also with increasing percentage of ethanol, followed 20
Content (mg/g)
15 p-Coumaric acid Chrysin Pinocembrin
10
Galangin
3.3. Method validation Good linearity was observed for phenolic acids and flavonoids used as propolis standards over the tested ranges (r2 > 0.999, see Table A in the supplementary material). The LOD value was in the range 0.5–0.8 g/ml for phenolic acids and 1.2–3.0 g/ml for flavonoids. The LOQ value was in the range 1.7–2.6 g/ml for phenolic acids and 3.7–9.9 g/ml for flavonoids. These LOD values were lower in comparison with the literature [4] and indicate that the proposed HPLC-DAD method has a good sensitivity for the determination of the phenolic composition of raw propolis. With regard to accuracy, the percentage recovery values were in the range 95.3–98.1% for phenolic acids and 94.1–101.3% for flavonoids (see Table B in the supplementary material). Considering the results of the recovery test, the developed method can be considered accurate. The low intra- and inter-day SD values for content (≤0.8 mg/g for chrysin and pinocembrin, see Table C in the supplementary material), and %RSD for retention times (≤0.3, see Table D in the supplementary material) and peak areas (≤2.2, see Table E in the supplementary material) indicate the high precision of both the extraction procedure and the chromatographic system. Regarding stability, reference standard solutions, subjected to HPLC analysis before and after MAE, did not show any appreciable change in the concentration after the microwave irradiation (Fig. 4) and degradation products were not detected. In addition, the chromatograms of propolis extracts analyzed every 12 h did not show changes in the profile of samples for 72 h. The validation data highlighted the suitability of the proposed method for the quali- and quantitative analysis of phenolic compounds in extracts obtained from raw propolis.
Pinobanks in-3-O-acetate Caffeic acid cinnamyl ester 5
0 Maceration
HRE
UAE
MAE
Fig. 3. Results obtained by the final comparison of the yields (mg/g) of six selected compounds from raw propolis (sample RP-1), using different extraction techniques. Compounds are listed in order of elution time.
3.4. Analysis of raw propolis samples The analysis of Italian samples of raw propolis was performed using the optimized MAE procedure and RP-HPLC method; the results are shown in Table 2. The overall chemical composition was the same in all the samples analyzed, including phenolic acids, flavones, flavonols, flavanones and dihydroflavonols. A significant variability in the
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Table 2 Content (mg/g) of phenolic acids and flavonoids in raw propolis samples (RP-1/RP-9) by HPLC-DAD.a Peak number
Compound name
RP-1
RP-2
RP-3
RP-4
RP-5
RP-6
RP-7
RP-8
RP-9
1 2 3 4 5
Caffeic acid p-Coumaric acid Ferulic acid Isoferulic acid 3,4-Dimethyl-caffeic acid (DMCA) Quercetin Cinnamic acid Pinobanksin-5-methyl-ether Quercetin-3-methyl-ether Apigenin Pinobanksin Kaempferol Isorhamnetin Luteolin-methyl-ether Quercetin-dimethyl-ether Cinnamylidene acetic acid Pinobanksin-5-methyl-ether3-O-acetate Quercetin-7-methyl-ether Quercetin-dimethyl-ether Caffeic acid prenyl ester Chrysin Caffeic acid prenyl ester Caffeic acid benzyl ester Pinocembrin Galangin Pinobanksin-3-O-acetate Caffeic acid phenylethyl ester (CAPE) Methoxy-chrysin p-Coumaric prenyl ester p-Coumaric benzyl ester Caffeic acid cinnamyl ester p-Coumaric prenyl ester Pinobanksin-3-O-propionate Pinobanksin-3-O-butyratec p-Coumaric cinnamyl ester Pinobanksin-3-O-pentanoatec Pinobanksin-3-O-hexanoatec p-Methoxy cinnamic acid cinnamyl ester Total phenolic acids Total flavones Total flavonols Total flavanones Total dihydroflavonols Total flavonoids Total phenolics
3.4 ± 0.1 8.6 ± 0.1 6.2 ± 0.1 1.8b 5.2 ± 0.2
2.7b 5.0 ± 0.1 4.6b 1.8b 5.6 ± 0.1
2.3 ± 0.2 3.2 ± 0.6 3.8 ± 0.9 1.3 ± 0.1 3.8 ± 0.2
3.7 ± 0.2 2.5 ± 0.1 2.5 ± 0.1 2.0b 6.6 ± 0.1
3.5 ± 0.2 7.9 ± 0.5 8.7 ± 0.5 1.4 ± 0.1 4.9 ± 0.2
6.7 ± 0.7 1.9 ± 0.1 2.3 ± 0.1 2.8 ± 0.2 12.1 ± 0.7
0.3b 0.3b 0.8b
6.1b 10.2 ± 0.6 8.3 ± 1.0 3.0 ± 0.4 10.6 ± 0.1
3.5 ± 0.1 1.0b 0.9b 1.8 ± 0.1 5.2 ± 0.1
0.9b 1.6 ± 0.1 4.0 ± 0.1 1.2b 1.0b 4.1 ± 0.2 1.2b 1.4b 0.8b 1.1b 6.9 ± 0.2
0.7 ± 0.1 1.5 ± 0.4 3.5 ± 0.3 0.9 ± 0.1 0.8b 3.7 ± 0.2 0.9 ± 0.1 0.9 ± 0.1 0.7 ± 0.1 0.7b 6.6 ± 0.3
0.5 ± 0.1 0.5 ± 0.2 0.7 ± 0.1 0.3b 0.4 ± 0.1 0.9 ± 0.1 0.4b 0.5 ± 0.1
1.6b 0.9 ± 0.1 7.2 ± 0.2 2.0b 1.3b 4.0 ± 0.1 2.2 ± 0.1 2.6 ± 0.1 1.4 ± 0.1 1.7 ± 0.1 6.1 ± 0.4
0.8b 0.6b 2.9 ± 0.1 0.8b 0.7b 2.0 ± 0.1 1.3 ± 0.1 1.0 ± 0.1 0.8b 0.7b 4.6 ± 0.2
2.5 ± 0.1 0.8 ± 0.3 9.4 ± 0.1 3.8 ± 0.2 2.0 ± 0.2 5.5 ± 0.4 2.5 ± 0.3 3.8 ± 0.3 2.4 ± 0.2 3.1 ± 0.2 5.4 ± 0.1 1.4 ± 0.1
1.5b 0.5 ± 0.1 3.5 ± 0.1 2.2 ± 0.1 3.0 ± 0.5 6.2 ± 0.2 2.1 ± 0.1 0.9 ± 0.1 2.4 ± 0.2 1.2 ± 0.1 1.0b 1.1b
1.1b 0.4b 3.0 ± 0.1 1.5 ± 0.1 1.8b 7.8b 1.8 ± 0.1 1.5 ± 0.1 1.6 ± 0.1 1.1 ± 0.1 0.9b
1.4b 2.0 ± 0.1 4.3 ± 0.1 14.2 ± 0.5 3.4 ± 0.1 10.0 ± 0.3 16.3 ± 0.5 8.1 ± 0.3 13.3 ± 0.6 6.6 ± 0.3
1.1 ± 0.1 1.4 ± 0.1 3.9 ± 0.1 13.0 ± 0.5 3.7b 9.6 ± 0.6 14.3 ± 1.4 6.8 ± 0.6 11.7 ± 0.8 6.0 ± 0.3
0.5 ± 0.1 0.4 ± 0.1 3.1 ± 0.2 4.2 ± 0.6 3.6 ± 0.3 3.3 ± 0.4 4.2 ± 0.7 1.1 ± 0.2 2.5 ± 0.4 2.3 ± 0.4
2.1 ± 0.1 2.8 ± 0.1 6.9 ± 0.1 18.5 ± 0.6 5.5 ± 0.1 13.6 ± 0.5 24.3 ± 0.8 12.3 ± 0.2 17.2 ± 1.1 7.1 ± 0.1
1.1 ± 0.1 1.3b 4.2 ± 0.2 9.9 ± 0.5 4.3 ± 0.2 11.3 ± 0.5 10.3 ± 0.5 6.3 ± 0.3 8.1 ± 0.4 4.9 ± 0.2
3.6 ± 0.2 6.1 ± 0.3 11.6 ± 0.9 33.0 ± 2.0 10.3 ± 0.8 22.3 ± 0.9 23.3 ± 1.1 16.3 ± 1.0 20.7 ± 1.6 13.1 ± 0.7
3.3 ± 0.1 4.4 ± 0.2 12.4 ± 2.2 23.4 ± 0.2 19.2 ± 0.3 17.2 ± 0.1 25.0 ± 0.4 12.5 ± 0.1 31.9 ± 0.2 9.6 ± 0.2
1.8b 2.6 ± 0.2 8.3 ± 0.3 16.8 ± 0.8 10.9 ± 0.3 7.7 ± 0.4 15.1 ± 0.6 9.3 ± 0.2 15.9 ± 0.8 4.8 ± 0.2
1.4b 0.3b 12.0 ± 0.2 16.3 ± 0.5 1.4b 2.2 ± 0.1 2.9 ± 0.1 1.6b 2.3 ± 0.1 1.9 ± 0.1 0.8b
1.2 ± 0.1 0.4b 6.2 ± 0.2 14.8 ± 0.6 0.8 ± 0.1 1.7 ± 0.3 2.5 ± 0.2 1.6 ± 0.1 1.8 ± 0.1 1.6 ± 0.2 1.0 ± 0.4
1.7 ± 0.1 0.6b 3.0 ± 0.1 17.2 ± 0.7 2.0 ± 0.1 3.1b 2.6 ± 0.2 2.1 ± 0.1 2.9 ± 0.1 2.7 ± 0.1 0.8 ± 0.1
1.2b 0.4b 9.1 ± 0.5 17.3 ± 0.9 0.8b 1.5 ± 0.1 1.5 ± 0.1 0.6 ± 0.1 1.5 ± 0.1 1.3 ± 0.1 0.5b
4.1 ± 0.2 0.7b 2.0b 24.8 ± 1.6 4.5 ± 0.3 6.9 ± 0.4 7.0 ± 0.5 1.5 ± 0.1 6.4 ± 0.5 4.2 ± 0.1 1.3 ± 0.3
4.0 ± 0.1 0.6b 8.8 ± 0.4 11.7 ± 0.3 2.1 ± 0.1 3.2 ± 0.2 4.5 ± 0.3 1.0 ± 0.2 2.6 ± 0.1 2.2 ± 0.1 0.7 ± 0.4
2.6 ± 0.1 0.6b 1.0b 7.0 ± 0.4 1.3 ± 0.1 1.9 ± 0.1 5.8 ± 1.0 0.6b 2.0 ± 0.1 1.2 ± 0.1 0.8 ± 0.1
81.3 ± 2.1 17.3 ± 0.6 17.2 ± 0.6 16.3 ± 0.5 30.7 ± 1.2 81.6 ± 2.9 172.2 ± 5.2
66.9 ± 2.0 15.8 ± 0.7 13.5 ± 1.0 14.3 ± 1.4 26.6 ± 2.0 70.1 ± 5.1 146.2 ± 7.4
36.4 ± 5.3 4.6 ± 0.7 4.1 ± 0.6 4.2 ± 0.7 6.6 ± 1.1 19.5 ± 3.1 61.5 ± 8.9
75.2 ± 1.8 22.9 ± 0.7 27.4 ± 0.5 24.3 ± 0.8 39.7 ± 1.0 114.3 ± 2.1 197.3 ± 4.2
79.4 ± 4.1 12.5 ± 0.6 13.4 ± 0.7 10.3 ± 0.5 18.7 ± 0.7 55.0 ± 2.4 140.1 ± 6.8
116.5 ± 6.9 41.5 ± 2.5 41.6 ± 2.7 23.3 ± 1.1 61.7 ± 3.6 168.0 ± 9.9 292.2 ± 16.7
5.0 ± 0.2 0.9b 0.3b 0.7b 0.6b 2.5b 7.9 ± 0.2
120.8 ± 2.4 32.8 ± 0.1 28.1 ± 0.5 25.0 ± 0.4 55.2 ± 0.2 141.1 ± 0.3 264.1 ± 2.8
54.7 ± 1.5 22.8 ± 0.9 20.7 ± 0.6 15.1 ± 0.6 37.5 ± 2.1 96.0 ± 3.9 152.8 ± 4.9
6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 – – – – – – –
Experimental conditions as in Section 2.2. a Data are expressed as mean (n = 6) ± SD. b SD < 0.05. c Or positional isomers.
content of total phenolics, including phenolic acids and flavonoids, was observed. The content of total phenolics ranged from 7.9 mg/g (sample RP-7) to 292.2 mg/g (sample RP-6). The amount of total phenolic acids ranged from 5.0 mg/g (sample RP-7) to 120.8 mg/g (RP-8) and the level of total flavonoids from 2.5 mg/g (sample RP7) to 168.0 mg/g (sample RP-6). The content of total flavonoids and total phenolic acids was in agreement with the literature [17]. On the basis of total flavonoid level, propolis with a content lower than 11% has been considered of low quality, while that with a content of 11–14%, 14–17% or >17% has been classified as acceptable, good and high quality, respectively [17]. In this context, samples RP-1, RP-2, RP-3, RP-5, RP-7 and RP-9 can be considered as low quality; sample RP-4 as acceptable; samples RP-6 and RP-8 as good. In the samples analyzed, the most abundant flavonoids were chrysin, pinocembrin, galangin and pinobanksin-3-O-acetate.
Regarding phenolic acids, caffeic acid, p-coumaric acid and ferulic acid were the most abundant ones. As for phenolic acid derivatives, 3,4-dimethyl caffeic acid (DMCA), caffeic acid prenyl, benzyl, phenylethyl (CAPE) and cinnamyl esters were the most representative compounds. 4. Conclusions An efficient closed-vessel MAE process was developed and optimized for the first time by means of a response surface experimental design methodology for the fast extraction of phenolic acids and flavonoids from raw propolis. Compared with traditional methods, the proposed MAE procedure allowed a reduction of extraction time and solvent use. The extraction efficiency of MAE was comparable with that obtained with conventional extraction methods and UAE, without degradation of the compounds of interest.
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The results of the HPLC analysis of real matrixes, carried out using a validated method based on the use of a fused-core C18 column, indicated a great variability in the content of the active compounds. In this context, the extraction procedure and HPLC method developed in this study can be considered reliable and useful tools for the comprehensive multi-component analysis of polyphenols in extracts of raw propolis to be used in apitherapy. Acknowledgments The authors are grateful to Kontak (Pozzo d’Adda, Milan, Italy) for the financial support. The authors acknowledge the “Fondazione Cassa di Risparmio di Modena” for funding the HPLC–ESI–MS system at the Centro Interdipartimentale Grandi Strumenti (CIGS) of the University of Modena and Reggio Emilia. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.jpba.2013.04.003. References [1] J.M. Sforcin, V. Bankova, Propolis: is there a potential for the development of new drugs? J. Ethnopharmacol. 133 (2011) 253–260. [2] A.M. Gómez-Caravaca, M. Gómez-Romero, D. Arráez-Román, A. SeguraCarretero, A. Fernández-Gutiérrez, Advances in the analysis of phenolic compounds in products derived from bees, J. Pharm. Biomed. Anal. 41 (2006) 1220–1234. [3] Y. Xu, L. Luo, B. Chen, Y. Fu, Recent developments of chemical components in propolis, Front. Biol. China 4 (2009) 385–391.
[4] F. Pellati, G. Orlandini, D. Pinetti, S. Benvenuti, HPLC-DAD and HPLC-ESI-MS/MS methods for metabolite profiling of propolis extracts, J. Pharm. Biomed. Anal. 55 (2011) 934–948. [5] P.K. Mukherjee, Extraction of herbal drugs, in: Quality Control of Herbal Drugs, Business Horizons, New Delhi, 2005, pp. 379–425. ´ [6] G. Romanik, E. Gilgenast, A. Przyjazny, M. Kaminski, Techniques of preparing plant material for chromatographic separation and analysis, J. Biochem. Biophys. Methods 70 (2007) 253–261. [7] H.F. Zhang, X.H. Yang, Y. Wang, Microwave assisted extraction of secondary metabolites from plants: current status and future directions, Trends Food Sci. Technol. 22 (2011) 672–688. [8] W. Routray, V. Orsat, Microwave-assisted extraction of flavonoids: a review, Food Bioprocess Technol. 5 (2012) 409–424. [9] B. Trusheva, D. Trunkova, V. Bankova, Different extraction methods of biologically active components from propolis: a preliminary study, Chem. Cent. J. 1 (2007) 13. [10] M. Biesaga, Influence of extraction methods on stability of flavonoids, J. Chromatogr. A 1218 (2011) 2505–2512. [11] Y.K. Park, M. Ikegaki, Preparation of water and ethanolic extracts of propolis and evaluation of the preparations, Biosci. Biotechnol. Biochem. 62 (1998) 2230–2232. [12] A. Liazid, M. Palma, J. Brighi, C.G. Barroso, Investigation on phenolic compounds stability during microwave-assisted extraction, J. Chromatogr. A 1140 (2007) 29–34. [13] International Conference on Harmonization of Technical Requirements for the Registration of Pharmaceuticals for Human Use (ICH), Guideline Q2(R1)Validation of Analytical Procedures: Text and Methodology, ICH Secretariat, c/o IFPMA, Geneva, 2005, pp. 1–13. [14] W. Xiao, L. Han, B. Shi, Microwave-assisted extraction of flavonoids from Radix astragali, Sep. Purif. Technol. 62 (2008) 614–618. [15] C.C. Teo, S.N. Tan, J.W. Hong Yong, C.S. Hew, E.S. Ong, Validation of green-solvent extraction combined with chromatographic chemical fingerprint to evaluate quality of Stevia rebaudiana Bertoni, J. Sep. Sci. 32 (2009) 613–622. [16] M.A. Bezerra, R.E. Santelli, E.P. Oliveira, L.S. Villar, L.A. Escaleira, Response surface methodology (RSM) as a tool for optimization in analytical chemistry, Talanta 76 (2008) 965–977. [17] C. Gardana, M. Scaglianti, P. Pietta, P. Simonetti, Analysis of the polyphenolic fraction of propolis from different sources by liquid chromatography–tandem mass spectrometry, J. Pharm. Biomed. Anal. 45 (2007) 390–399.
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An efficient chemical analysis of phenolic acids and flavonoids in raw propolis by microwave-assisted extraction combined with high-performance liquid chromatography using the fused-core technology Federica Pellati ∗ , Francesco Pio Prencipe, Davide Bertelli, Stefania Benvenuti Department of Life Sciences, University of Modena and Reggio Emilia, Via G. Campi 183, 41125 Modena, Italy
a r t i c l e
i n f o
Article history: Received 8 February 2013 Received in revised form 3 April 2013 Accepted 5 April 2013 Available online xxx Keywords: Propolis Flavonoids Phenolic acids Microwave-assisted extraction Fused-core HPLC
a b s t r a c t A closed-vessel microwave-assisted extraction (MAE) technique was optimized for the first time for the extraction of polyphenols from raw propolis. The results obtained by means of response surface experimental design methodology showed that the best global response was reached when the extraction temperature was set at 106 ◦ C, the solvent composition close to EtOH–H2 O 80:20 (v/v), with an extraction time of 15 min. In comparison with other techniques, such as maceration, heat reflux extraction (HRE) and ultrasound-assisted extraction (UAE), the extraction with MAE was improved by shorter extraction time and lower volume of solvent needed. The HPLC analyses of propolis extracts were carried out on a fused-core Ascentis Express C18 column (150 mm × 3.0 mm I.D., 2.7 m), with a gradient mobile phase composed by 0.1% formic acid in water and acetonitrile. Detection was performed by DAD and MS. The method validation indicated that the correlation coefficients were >0.999; the limit of detection was in the range 0.5–0.8 g/ml for phenolic acids and 1.2–3.0 g/ml for flavonoids; the recovery range was 95.3–98.1% for phenolic acids and 94.1–101.3% for flavonoids; the intra- and inter-day %RSD values for retention times and peak areas were ≤0.3 and 2.2%, respectively. The quali- and quantitative analysis of polyphenols in Italian samples of raw propolis was performed with the validated method. Total phenolic acids ranged from 5.0 to 120.8 mg/g and total flavonoids from 2.5 to 168.0 mg/g. The proposed MAE procedure and HPLC method can be considered reliable and useful tools for the comprehensive multi-component analysis of polyphenols in propolis extracts to be used in apitherapy. © 2013 Elsevier B.V. All rights reserved.
1. Introduction Propolis is a resinous material collected by honeybees (Apis mellifera L.) from several tree species [1]. In regions with temperate climate, the resin is collected mainly from the buds and cracks in the bark of Poplar trees [2]. Propolis is characterized by a series of biological properties, such as antibacterial, antiviral, antifungal, anti-inflammatory, antioxidant, antiproliferative, immunostimulating [1]. Typical applications of propolis include herbal products for cold syndrome and dermatological preparations [3]. Propolis extracts are also used to prevent and treat oral inflammations [3].
Abbreviations: MAE, microwave-assisted extraction; HRE, heat reflux extraction; UAE, ultrasound-assisted extraction; CCD, central composite design; DMCA, 3,4dimethyl caffeic acid; CAPE, caffeic acid phenylethyl ester. ∗ Corresponding author. Tel.: +39 059 205 5144; fax: +39 059 205 5131. E-mail address: [email protected] (F. Pellati). 0731-7085/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.jpba.2013.04.003
The chemical composition of propolis is known to be very complex and influenced by the geographic origin [3]. In propolis from temperate zones, the most representative biologically active compounds are polyphenols, including phenolic acids and flavonoids [2,3]. Raw propolis cannot be used as crude material, but it must be purified by solvent extraction to remove the useless material and preserve the active polyphenolic fraction [2,3]. Hydroalcoholic solvents have been described as the most suitable media for the extraction of biologically active phenolic components from raw propolis [2]. Regarding the extraction techniques used for the chemical analysis of raw propolis, maceration at room temperature and heat reflux extraction (HRE) have been widely applied [2,4]. However, conventional extraction methods are often timeconsuming and require high solvent volumes, raising process costs and reducing environmental sustainability [5,6]. For this reason, the exploration of innovative extraction techniques that could solve some of the above mentioned problems has been considered [6]. In this ambit, microwave-assisted extraction (MAE) represents a
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reliable alternative to traditional extraction techniques [7] and it has been widely applied to the extraction of flavonoids from plant material [8]. Several examples in the ambit of phytochemical analysis suggested that MAE has some considerable merits, such as shorter extraction time, higher extraction yield and less solvent consumption compared to conventional extraction methods [7]. Different types of MAE systems can be applied to the extraction of phenolics from natural products, including microwave ovens operating at the atmospheric pressure and closed-vessel equipments under controlled temperature and pressure [7,8]. In a previous study, Trusheva et al. [9] used a multimodal household microwave oven at 800 W for the extraction of polyphenols from propolis; the authors have observed the presence of unwanted compounds in the extracts and a decrease in the amount of polyphenols, probably due to the very high microwave power applied [10]. In addition, the use of domestic ovens without technical modifications should be avoided, because of the low safety, reliability and reproducibility which characterize these equipments [7]. In this study, a response surface experimental design methodology coupled with central composite design (CCD) was applied for the first time to optimize the extraction of polyphenols from raw propolis using MAE. To the best of our knowledge, closed-vessel MAE with a system operating at controlled temperature and low irradiation power has never been applied before to the extraction of phenolic compounds from raw propolis. The extraction efficiency of MAE was compared with that of other techniques previously described in the literature [2,4], including maceration, HRE and ultrasound-assisted extraction (UAE), in terms of yield, time and solvent usage. The analysis of polyphenols in propolis extracts was carried out by an improved RP-HPLC method using a fused-core C18 column. The method was fully validated and its practical applicability was demonstrated by the analysis of Italian samples of raw propolis to provide a complete chemical analysis of their biologically active constituents.
2. Materials and methods 2.1. Propolis samples, chemicals and solvents Nine samples of raw propolis (indicated in the text as RP-1/RP-9) were collected from Apis mellifera hives located in different Italian regions in Spring 2012 and stored at −20 ◦ C until chemical analysis. The frozen samples were finely powdered using a mortar and a pestle before the extraction procedure. Sample labeled as RP-1 was used for the optimization of the extraction conditions, because it was available in higher amount. Caffeic acid, p-coumaric acid, ferulic acid, quercetin, pinocembrin, cinnamic acid, apigenin, kaempferol and galangin were from Sigma–Aldrich–Fluka (Milan, Italy). Isorhamnetin and luteolin were from Roth (Karlsruhe, Germany). Chrysin was from Extrasynthese (Genay, France). The purity of standard compounds was always higher than 97%, as confirmed by HPLC-DAD analysis. HPLC-grade acetonitrile (ACN), formic acid and analytical grade absolute ethanol (EtOH) were from Sigma (Milan, Italy). Water was purified using a Milli-Q Plus185 system from Millipore (Milford, MA, USA).
2.2. HPLC-DAD conditions HPLC analyses were performed on an Agilent Technologies (Waldbronn, Germany) modular model 1100 system consisting of a vacuum degasser, a quaternary pump, an autosampler, a thermostatted column compartment and a diode array detector (DAD).
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The chromatograms were recorded using an Agilent Chemstation for LC and LC–MS systems (Rev. B.01.03). An Ascentis Express C18 column (150 mm × 3.0 mm I.D., 2.7 m, Supelco, Bellefonte, PA, USA) was used for the HPLC analysis. The mobile phase was composed by 0.1% formic acid in H2 O (A) and ACN (B). The gradient elution was modified as follows: 0–3 min 20% B, 3–10 min from 20 to 30% B, 10–40 min from 30 to 40% B, 40–50 min from 40 to 60% B, 50–60 min from 60 to 80% B, 60–65 min from 80 to 50% B. The post-running time was 10 min. The flow rate was 0.6 ml/min. The column temperature was set at 30 ◦ C. The sample injection volume was 2 l. The DAD acquisitions were carried out in the range 190–450 nm and chromatograms were integrated at 265 nm (for chrysin and galangin), 290 nm (for cinnamic acid and pinocembrin), 320 nm (for caffeic acid, p-coumaric acid and ferulic acid), 338 nm (for apigenin and luteolin) and 370 nm (for quercetin, isorhamnetin and kaempferol). Three injections were performed for each sample. 2.3. HPLC–ESI–MS and MS2 conditions HPLC-ESI-MS and MS2 analyses were carried out using an Agilent Technologies modular 1200 system, equipped with a vacuum degasser, a binary pump, an autosampler, a thermostated column compartment and a 6310A ion trap mass analyzer with an ESI ion source. The HPLC column and the applied chromatographic conditions were the same as reported for the HPLC-DAD system. The flow rate was split 3:1 before the ESI source. The experimental parameters were set as follows: the capillary voltage was 3.5 kV, the nebulizer (N2 ) pressure was 32 psi, the drying gas temperature was 350 ◦ C, the drying gas flow was 11 l/min and the skimmer voltage was 40 V. Data were acquired by Agilent 6300 Series Ion Trap LC/MS system software (version 6.2). The mass spectrometer was operated in the full-scan positive and negative ion modes in the m/z range 100–1000. MS2 spectra were automatically performed with helium as the collision gas by using the SmartFrag function. 2.4. Methods for the extraction of phenolics from raw propolis Four methods were applied and compared in order to obtain a high extraction efficiency of phenolic components from raw propolis. In each case, the applied sample-to-solvent ratio was 1:10 (w/v), as previously described in the literature [2,9]. The extraction procedure was repeated twice for each method. 2.4.1. Maceration, heat reflux extraction (HRE) and ultrasound-assisted extraction (UAE) Maceration extraction was performed on a weighed amount of sample (1.00 g) with 10 ml of EtOH–H2 O (80:20, v/v) at room temperature for 24 h under stirring. The HRE was carried out by refluxing a weighed amount of sample (1.00 g) with 10 ml of EtOH–H2 O (80:20, v/v) at 70 ◦ C for 1 h under stirring using a water bath. The UAE was performed on a weighed amount of sample (1.00 g) with 10 ml of EtOH–H2 O (80:20, v/v) at 70 ◦ C for 1 h using an ultrasonic bath (Sonorex RK-100H, Bandelin, Berlin, Germany). At the end of each of these extraction procedures, the mixture was centrifuged at 4000 rpm for 5 min. The supernatant solution was then paper filtered in a vacuum into a 10 ml volumetric flask and the solvent was added to the final volume. One ml aliquot of each propolis hydroalcoholic extract was diluted 1:5 (v/v) with the extraction solvent in a volumetric flask, filtered through a 0.45 m PTFE filter into a HPLC vial and injected into the HPLC system. Three injections were performed for each sample.
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Table 1 Factor levels and experimental domain applied to optimize the MAE experimental conditions. Factor
Solvent composition (EtOH–H2 O ratio, v/v) Temperature (T, ◦ C) Time (t, min)
Solvent composition (EtOH–H2 O ratio, v/v) Temperature (T, ◦ C) a b
First experimental domain −˛a
−1
0
1
˛a
49.7 66.4 6.6
60.0 80.0 10.0
75.0 100.0 15.0
90.0 120.0 20.0
100.2 133.6 23.4
Second experimental domain −˛b −1
0
1
˛b
53.8 71.7
75.0 100.0
90.0 120.0
96.2 128.3
60.0 80.0
˛ = 1.682. ˛ = 1.414.
2.4.2. Microwave-assisted extraction (MAE) Regarding MAE, a weighed amount of ground sample (0.50 g) was extracted with 5 ml of solvent in a 10 ml glass vessel using a monomode focused microwave apparatus with a closed-vessel system (Discover instrument, CEM, Metthews, NC, USA). The microwave equipment used in this study allowed extraction temperature, pressure and time to be programmed. Microwave power, with a maximum of 300 W and a magnetron frequency of 2450 MHz, and pressure were dynamically adjusted by temperature and power feedback control. The instrument was able to provide continuous non-pulse microwave heating, focusing on a single cavity where the sample was placed. During the extraction, magnetic stirring was applied to homogenize the sample. The optimized experimental conditions were set as follows: extraction solvent EtOH–H2 O (80:20, v/v), extraction temperature at 106 ◦ C and extraction time of 15 min. After the extraction time had elapsed, the vessel was allowed to cool at room temperature before opening. After centrifugation for 5 min at 4000 rpm, the supernatant solution was paper filtered in a vacuum into a 5 ml volumetric flask and the solvent was added to the final volume. Further processing of the samples prior to HPLC analysis was as described above.
2.5. Experimental design In order to optimize the extraction parameters of phenolics from raw propolis using MAE, an experimental design was applied. The optimization of MAE conditions was performed using a CCD (with ˛ = 1.682), which was based on a 23 factorial design (each experiment performed in triplicate) plus six axial points (each experiment performed in triplicate) plus six replicates in the center of the design. For MAE optimization, the variables chosen were the solvent composition (EtOH–H2 O ratio, v/v), the extraction temperature (T, ◦ C) and the extraction time (t, min). The factor levels and experimental domain are shown in Table 1. In particular, the solvent composition varied from EtOH–H2 O 50:50 (v/v) to 100% EtOH, covering the concentration range characterized by a high extraction yield for polyphenols from propolis [11]. The extraction temperature varied from 66 to 134 ◦ C, avoiding higher temperature that could cause the degradation of flavonoids, as described by Liazid et al. [12]. Finally, the extraction time varied from 7 to 23 min in order to obtain a fast extraction method. A total of forty-eight experiments were performed in randomized order. In view of the results of these experiments, the two most significant variables (EtOH–H2 O volume ratio and temperature) were selected to perform a second CCD with ˛ = 1.414, based on a 22 factorial design (each experiment performed in duplicate) plus four axial points (each experiment performed in duplicate) plus four replicates in the center of the design. The experimental domain was defined taking into account the results obtained from the first
CCD with some adjustments (Table 1), i.e. the solvent composition varied from EtOH–H2 O 54:46 (v/v) to 96% EtOH and the extraction temperature from 72 to 128 ◦ C. The extraction time was kept constant at 15 min. Twenty experiments were performed in randomized order. The analysis of the second CCD was performed by response surface methodology; this surface is approximated by a polynomial function, using a quadratic model, and represents a good description of the relationship between the experimental variables and the response within a limited experimental domain. Statistical analysis was performed using Statistica v. 6.1 (Statsoft Inc., Tulsa, OK, USA).
2.6. HPLC-DAD method validation The validation of the HPLC-DAD method was performed to show compliance with international requirements for analytical techniques for the quality control of pharmaceuticals (ICH guidelines) [13]. Regarding linearity, the stock solution of each compound commercially available (caffeic acid, p-coumaric acid, ferulic acid, quercetin, cinnamic acid, apigenin, kaempferol, isorhamnetin, luteolin, chrysin, pinocembrin and galangin) was prepared as follows: an accurately weighed amount of each pure standard compound (2–6 mg) was placed into a 10 ml volumetric flask; EtOH was added and the solution was diluted to volume with the same solvent. The external standard calibration curve was generated using seven data points. Injections were performed in triplicate for each concentration level. The calibration curve was obtained by plotting the peak area of the compound at each level versus the concentration of the sample. For each compound, the limit of detection (LOD) and the limit of quantification (LOQ) were experimentally verified by HPLC analysis of serial dilutions of a standard solution to reach a signal-to-noise (S/N) ratio of 3 and 10, respectively. The accuracy of the analytical procedure was evaluated using the recovery test. This involved the addition of a known quantity of standard compound to half the sample weight of raw propolis (sample RP-1) to reach 100% of the test concentration. The fortified samples were then extracted using the optimized MAE conditions and analyzed by the proposed HPLC method. The precision of the extraction technique was validated by repeating the extraction procedure on the same sample of raw propolis (RP-1) at the best MAE conditions. An aliquot of each extract was then injected and quantified. The extraction was performed in duplicate on three different days with newly prepared mobile phase and samples. The precision of the chromatographic system was tested by performing intra- and inter-day multiple injections of a hydroalcoholic extract of propolis (RP-1) and then checking the %RSD of retention times and peak areas. Six injections were performed each day for three consecutive days.
F. Pellati et al. / Journal of Pharmaceutical and Biomedical Analysis 81–82 (2013) 126–132 mAU 700
129
1 21
600 24
5
500
7+8
400
23
300
26
11
2 4
31
200
36
20
16
22
25
27
3
34+35 9
100 6
10 1314 12 15
17
32
28
18
37
33
19 29
38
30
0 0
10
20
30
40
50
min
Fig. 1. Chromatogram obtained by RP–HPLC–DAD analysis of a representative propolis extract (sample RP-6) at 290 nm on the fused-core column. Experimental conditions as in Section 2.2. For peak identification, see Table 2.
The stability of the analytes during MAE was evaluated with reference standard solutions subjected to HPLC analysis before and after the extraction treatment under the optimized conditions. Stability was also tested using propolis extracts stored in amber glass flasks at 4 ◦ C and at room temperature (about 25 ◦ C) and analyzed every 12 h. 3. Results and discussion 3.1. HPLC method development and optimization An initial comparison of fully porous and fused-core C18 columns for the HPLC analysis of polyphenols in raw propolis was carried out. In particular, a method initially developed on a fully porous Ascentis C18 column for the analysis of commercially available propolis extracts [4] was transferred to a fused-core Ascentis Express C18 column. A parallel careful adaptation of the experimental conditions was performed. In this way, the total analysis time was decreased from 92 to 65 min, with a consequent advantage in terms of time and solvent usage. Under these conditions, the highest observed backpressure was about 300 bar, within the pressure limit of conventional HPLC systems. The HPLC-DAD analysis at 290 nm of a hydroalcoholic extract obtained from a representative sample of raw propolis (RP-6) indicated a very complex composition, as shown in Fig. 1. A total of 38 compounds were identified in propolis samples, on the basis of their UV, MS and MS2 data, which were compared with those of the reference standards and with the literature [4]. Taking into account the complexity of the sample, the chromatographic separation can be considered satisfactory. 3.2. Optimization of extraction conditions Four methods were evaluated in order to obtain an efficient and rapid extraction of polyphenols from raw propolis. The comparison was carried out between conventional extraction methods (including maceration, HRE and UAE) [2,4] and MAE, which represents a more innovative extraction procedure. Since closed-vessel MAE has never been investigated for the extraction of phenolics from raw propolis, its experimental parameters were optimized. For a closed-vessel MAE system, the variables that could affect the extraction efficiency include temperature, solvent composition, time, microwave power, pressure and sample-to-solvent ratio [8,14,15]. In this study, the attention was focused on temperature, solvent composition and time, which are
recognized as representative parameters affecting the efficiency of MAE [8]. It should be noted that the microwave power was not chosen for the optimization of the extraction conditions, because the closed-vessel MAE system used in this work regulates the microwave power depending on the selected temperature. Furthermore, in view of the solvent mixtures used in this study, the microwave system reached the desired temperature at low irradiation power: in fact, under the applied conditions, the microwave power never exceeded 120 W, thus avoiding a possible degradation of flavonoids due to high irradiation energy [10]. Pressure is another important experimental variable for closed-vessel MAE, but it is directly dependent on temperature [15] and, therefore, it was not chosen for method optimization. Other parameters, such as sample amount and solvent volume were kept constant, with a 1:10 (w/v) ratio [2,9]. Response surface experimental design methodology coupled with CCD is a useful tool to optimize procedures depending on several variables [16] and, therefore, it was applied in this study. A response based on the total amount (mg/g) of the most representative propolis constituents under the conditions of the design was studied. The experimental values of the factors chosen for the optimization of MAE parameters were selected in order to cover a wide range of conditions. The data obtained were evaluated by ANOVA, the level of significance being set at 5%, to evaluate the statistical significance of each factor and interactions between the different factors. The quadratic term of the solvent composition was the most significant parameter (at p < 0.05) on the extraction yield of the compounds of interest. The linear and the quadratic term of the extraction time, as well as its interaction with the other factors, did not show a significant influence on the efficiency of the extraction process. A second CCD was thus set up, focusing on the solvent composition and the extraction temperature. As a confirmation of what observed with the previous CCD, the linear and the quadratic term of the solvent composition were the most significant parameters (at p < 0.05), having a strong influence on the extraction yield of phenolics from raw propolis. The interaction between solvent composition and extraction temperature was also found to be significant in the extraction procedure. Fig. 2 shows the response surface plot for the total amount of the extracted compounds versus solvent composition and extraction temperature, with an extraction time equal to 15 min. The experimental results showed that the best global response, within the range studied, was reached when the extraction temperature was set at 106 ◦ C, the extraction solvent close to EtOH–H2 O
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Concentration ( µg/mL)
400
300
Before MAE
200
After MAE 100
Galangin
Pinocembrin
Chrysin
Isorhamnetin
Kaempferol
Apigenin
Cinnamic acid
Quercetin
Ferulic acid
p-Coumaric acid
Caffeic acid
0
Fig. 4. Stability study of phenolic acids and flavonoids used as propolis standards subjected to optimized MAE conditions.
by HPLC analysis. The results indicated that the amount of polyphenols obtained from the residue after the first extraction was below the LOQ values in all cases. These results indicated that MAE represents an attractive sample preparation method and has good potential for the extraction of polyphenols from raw propolis. Fig. 2. Response surface plot for total amount (mg/g) of six representative polyphenolic compounds extracted from raw propolis versus solvent composition (EtOH–H2 O ratio, v/v) and extraction temperature (T, ◦ C), with a constant extraction time (t) equal to 15 min.
80:20 (v/v), with an extraction time of 15 min. These values were therefore selected for the extraction of polyphenols from raw propolis in all the subsequent analyses. The total amount of representative propolis phenolic compounds extracted under optimized MAE conditions was then compared with those obtained with maceration, HRE and UAE. As shown in Fig. 3, the MAE technique allowed to obtain the yield of other methods in only 15 min. With the assistance of the fast heating of microwave irradiation, the extraction with MAE was improved by a reduced extraction time and half volume of solvent needed. The composition of propolis extracts processed by closed-vessel MAE was monitored in detail by HPLC-DAD at five different wavelengths, by MS and MS2 , and the results indicated the absence of unwanted compounds. In addition, degradation products were not observed in the HPLC chromatograms. An investigation on the residue composition after MAE was also performed, by carrying out further extraction steps with the same solvents and also with increasing percentage of ethanol, followed 20
Content (mg/g)
15 p-Coumaric acid Chrysin Pinocembrin
10
Galangin
3.3. Method validation Good linearity was observed for phenolic acids and flavonoids used as propolis standards over the tested ranges (r2 > 0.999, see Table A in the supplementary material). The LOD value was in the range 0.5–0.8 g/ml for phenolic acids and 1.2–3.0 g/ml for flavonoids. The LOQ value was in the range 1.7–2.6 g/ml for phenolic acids and 3.7–9.9 g/ml for flavonoids. These LOD values were lower in comparison with the literature [4] and indicate that the proposed HPLC-DAD method has a good sensitivity for the determination of the phenolic composition of raw propolis. With regard to accuracy, the percentage recovery values were in the range 95.3–98.1% for phenolic acids and 94.1–101.3% for flavonoids (see Table B in the supplementary material). Considering the results of the recovery test, the developed method can be considered accurate. The low intra- and inter-day SD values for content (≤0.8 mg/g for chrysin and pinocembrin, see Table C in the supplementary material), and %RSD for retention times (≤0.3, see Table D in the supplementary material) and peak areas (≤2.2, see Table E in the supplementary material) indicate the high precision of both the extraction procedure and the chromatographic system. Regarding stability, reference standard solutions, subjected to HPLC analysis before and after MAE, did not show any appreciable change in the concentration after the microwave irradiation (Fig. 4) and degradation products were not detected. In addition, the chromatograms of propolis extracts analyzed every 12 h did not show changes in the profile of samples for 72 h. The validation data highlighted the suitability of the proposed method for the quali- and quantitative analysis of phenolic compounds in extracts obtained from raw propolis.
Pinobanks in-3-O-acetate Caffeic acid cinnamyl ester 5
0 Maceration
HRE
UAE
MAE
Fig. 3. Results obtained by the final comparison of the yields (mg/g) of six selected compounds from raw propolis (sample RP-1), using different extraction techniques. Compounds are listed in order of elution time.
3.4. Analysis of raw propolis samples The analysis of Italian samples of raw propolis was performed using the optimized MAE procedure and RP-HPLC method; the results are shown in Table 2. The overall chemical composition was the same in all the samples analyzed, including phenolic acids, flavones, flavonols, flavanones and dihydroflavonols. A significant variability in the
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Table 2 Content (mg/g) of phenolic acids and flavonoids in raw propolis samples (RP-1/RP-9) by HPLC-DAD.a Peak number
Compound name
RP-1
RP-2
RP-3
RP-4
RP-5
RP-6
RP-7
RP-8
RP-9
1 2 3 4 5
Caffeic acid p-Coumaric acid Ferulic acid Isoferulic acid 3,4-Dimethyl-caffeic acid (DMCA) Quercetin Cinnamic acid Pinobanksin-5-methyl-ether Quercetin-3-methyl-ether Apigenin Pinobanksin Kaempferol Isorhamnetin Luteolin-methyl-ether Quercetin-dimethyl-ether Cinnamylidene acetic acid Pinobanksin-5-methyl-ether3-O-acetate Quercetin-7-methyl-ether Quercetin-dimethyl-ether Caffeic acid prenyl ester Chrysin Caffeic acid prenyl ester Caffeic acid benzyl ester Pinocembrin Galangin Pinobanksin-3-O-acetate Caffeic acid phenylethyl ester (CAPE) Methoxy-chrysin p-Coumaric prenyl ester p-Coumaric benzyl ester Caffeic acid cinnamyl ester p-Coumaric prenyl ester Pinobanksin-3-O-propionate Pinobanksin-3-O-butyratec p-Coumaric cinnamyl ester Pinobanksin-3-O-pentanoatec Pinobanksin-3-O-hexanoatec p-Methoxy cinnamic acid cinnamyl ester Total phenolic acids Total flavones Total flavonols Total flavanones Total dihydroflavonols Total flavonoids Total phenolics
3.4 ± 0.1 8.6 ± 0.1 6.2 ± 0.1 1.8b 5.2 ± 0.2
2.7b 5.0 ± 0.1 4.6b 1.8b 5.6 ± 0.1
2.3 ± 0.2 3.2 ± 0.6 3.8 ± 0.9 1.3 ± 0.1 3.8 ± 0.2
3.7 ± 0.2 2.5 ± 0.1 2.5 ± 0.1 2.0b 6.6 ± 0.1
3.5 ± 0.2 7.9 ± 0.5 8.7 ± 0.5 1.4 ± 0.1 4.9 ± 0.2
6.7 ± 0.7 1.9 ± 0.1 2.3 ± 0.1 2.8 ± 0.2 12.1 ± 0.7
0.3b 0.3b 0.8b
6.1b 10.2 ± 0.6 8.3 ± 1.0 3.0 ± 0.4 10.6 ± 0.1
3.5 ± 0.1 1.0b 0.9b 1.8 ± 0.1 5.2 ± 0.1
0.9b 1.6 ± 0.1 4.0 ± 0.1 1.2b 1.0b 4.1 ± 0.2 1.2b 1.4b 0.8b 1.1b 6.9 ± 0.2
0.7 ± 0.1 1.5 ± 0.4 3.5 ± 0.3 0.9 ± 0.1 0.8b 3.7 ± 0.2 0.9 ± 0.1 0.9 ± 0.1 0.7 ± 0.1 0.7b 6.6 ± 0.3
0.5 ± 0.1 0.5 ± 0.2 0.7 ± 0.1 0.3b 0.4 ± 0.1 0.9 ± 0.1 0.4b 0.5 ± 0.1
1.6b 0.9 ± 0.1 7.2 ± 0.2 2.0b 1.3b 4.0 ± 0.1 2.2 ± 0.1 2.6 ± 0.1 1.4 ± 0.1 1.7 ± 0.1 6.1 ± 0.4
0.8b 0.6b 2.9 ± 0.1 0.8b 0.7b 2.0 ± 0.1 1.3 ± 0.1 1.0 ± 0.1 0.8b 0.7b 4.6 ± 0.2
2.5 ± 0.1 0.8 ± 0.3 9.4 ± 0.1 3.8 ± 0.2 2.0 ± 0.2 5.5 ± 0.4 2.5 ± 0.3 3.8 ± 0.3 2.4 ± 0.2 3.1 ± 0.2 5.4 ± 0.1 1.4 ± 0.1
1.5b 0.5 ± 0.1 3.5 ± 0.1 2.2 ± 0.1 3.0 ± 0.5 6.2 ± 0.2 2.1 ± 0.1 0.9 ± 0.1 2.4 ± 0.2 1.2 ± 0.1 1.0b 1.1b
1.1b 0.4b 3.0 ± 0.1 1.5 ± 0.1 1.8b 7.8b 1.8 ± 0.1 1.5 ± 0.1 1.6 ± 0.1 1.1 ± 0.1 0.9b
1.4b 2.0 ± 0.1 4.3 ± 0.1 14.2 ± 0.5 3.4 ± 0.1 10.0 ± 0.3 16.3 ± 0.5 8.1 ± 0.3 13.3 ± 0.6 6.6 ± 0.3
1.1 ± 0.1 1.4 ± 0.1 3.9 ± 0.1 13.0 ± 0.5 3.7b 9.6 ± 0.6 14.3 ± 1.4 6.8 ± 0.6 11.7 ± 0.8 6.0 ± 0.3
0.5 ± 0.1 0.4 ± 0.1 3.1 ± 0.2 4.2 ± 0.6 3.6 ± 0.3 3.3 ± 0.4 4.2 ± 0.7 1.1 ± 0.2 2.5 ± 0.4 2.3 ± 0.4
2.1 ± 0.1 2.8 ± 0.1 6.9 ± 0.1 18.5 ± 0.6 5.5 ± 0.1 13.6 ± 0.5 24.3 ± 0.8 12.3 ± 0.2 17.2 ± 1.1 7.1 ± 0.1
1.1 ± 0.1 1.3b 4.2 ± 0.2 9.9 ± 0.5 4.3 ± 0.2 11.3 ± 0.5 10.3 ± 0.5 6.3 ± 0.3 8.1 ± 0.4 4.9 ± 0.2
3.6 ± 0.2 6.1 ± 0.3 11.6 ± 0.9 33.0 ± 2.0 10.3 ± 0.8 22.3 ± 0.9 23.3 ± 1.1 16.3 ± 1.0 20.7 ± 1.6 13.1 ± 0.7
3.3 ± 0.1 4.4 ± 0.2 12.4 ± 2.2 23.4 ± 0.2 19.2 ± 0.3 17.2 ± 0.1 25.0 ± 0.4 12.5 ± 0.1 31.9 ± 0.2 9.6 ± 0.2
1.8b 2.6 ± 0.2 8.3 ± 0.3 16.8 ± 0.8 10.9 ± 0.3 7.7 ± 0.4 15.1 ± 0.6 9.3 ± 0.2 15.9 ± 0.8 4.8 ± 0.2
1.4b 0.3b 12.0 ± 0.2 16.3 ± 0.5 1.4b 2.2 ± 0.1 2.9 ± 0.1 1.6b 2.3 ± 0.1 1.9 ± 0.1 0.8b
1.2 ± 0.1 0.4b 6.2 ± 0.2 14.8 ± 0.6 0.8 ± 0.1 1.7 ± 0.3 2.5 ± 0.2 1.6 ± 0.1 1.8 ± 0.1 1.6 ± 0.2 1.0 ± 0.4
1.7 ± 0.1 0.6b 3.0 ± 0.1 17.2 ± 0.7 2.0 ± 0.1 3.1b 2.6 ± 0.2 2.1 ± 0.1 2.9 ± 0.1 2.7 ± 0.1 0.8 ± 0.1
1.2b 0.4b 9.1 ± 0.5 17.3 ± 0.9 0.8b 1.5 ± 0.1 1.5 ± 0.1 0.6 ± 0.1 1.5 ± 0.1 1.3 ± 0.1 0.5b
4.1 ± 0.2 0.7b 2.0b 24.8 ± 1.6 4.5 ± 0.3 6.9 ± 0.4 7.0 ± 0.5 1.5 ± 0.1 6.4 ± 0.5 4.2 ± 0.1 1.3 ± 0.3
4.0 ± 0.1 0.6b 8.8 ± 0.4 11.7 ± 0.3 2.1 ± 0.1 3.2 ± 0.2 4.5 ± 0.3 1.0 ± 0.2 2.6 ± 0.1 2.2 ± 0.1 0.7 ± 0.4
2.6 ± 0.1 0.6b 1.0b 7.0 ± 0.4 1.3 ± 0.1 1.9 ± 0.1 5.8 ± 1.0 0.6b 2.0 ± 0.1 1.2 ± 0.1 0.8 ± 0.1
81.3 ± 2.1 17.3 ± 0.6 17.2 ± 0.6 16.3 ± 0.5 30.7 ± 1.2 81.6 ± 2.9 172.2 ± 5.2
66.9 ± 2.0 15.8 ± 0.7 13.5 ± 1.0 14.3 ± 1.4 26.6 ± 2.0 70.1 ± 5.1 146.2 ± 7.4
36.4 ± 5.3 4.6 ± 0.7 4.1 ± 0.6 4.2 ± 0.7 6.6 ± 1.1 19.5 ± 3.1 61.5 ± 8.9
75.2 ± 1.8 22.9 ± 0.7 27.4 ± 0.5 24.3 ± 0.8 39.7 ± 1.0 114.3 ± 2.1 197.3 ± 4.2
79.4 ± 4.1 12.5 ± 0.6 13.4 ± 0.7 10.3 ± 0.5 18.7 ± 0.7 55.0 ± 2.4 140.1 ± 6.8
116.5 ± 6.9 41.5 ± 2.5 41.6 ± 2.7 23.3 ± 1.1 61.7 ± 3.6 168.0 ± 9.9 292.2 ± 16.7
5.0 ± 0.2 0.9b 0.3b 0.7b 0.6b 2.5b 7.9 ± 0.2
120.8 ± 2.4 32.8 ± 0.1 28.1 ± 0.5 25.0 ± 0.4 55.2 ± 0.2 141.1 ± 0.3 264.1 ± 2.8
54.7 ± 1.5 22.8 ± 0.9 20.7 ± 0.6 15.1 ± 0.6 37.5 ± 2.1 96.0 ± 3.9 152.8 ± 4.9
6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 – – – – – – –
Experimental conditions as in Section 2.2. a Data are expressed as mean (n = 6) ± SD. b SD < 0.05. c Or positional isomers.
content of total phenolics, including phenolic acids and flavonoids, was observed. The content of total phenolics ranged from 7.9 mg/g (sample RP-7) to 292.2 mg/g (sample RP-6). The amount of total phenolic acids ranged from 5.0 mg/g (sample RP-7) to 120.8 mg/g (RP-8) and the level of total flavonoids from 2.5 mg/g (sample RP7) to 168.0 mg/g (sample RP-6). The content of total flavonoids and total phenolic acids was in agreement with the literature [17]. On the basis of total flavonoid level, propolis with a content lower than 11% has been considered of low quality, while that with a content of 11–14%, 14–17% or >17% has been classified as acceptable, good and high quality, respectively [17]. In this context, samples RP-1, RP-2, RP-3, RP-5, RP-7 and RP-9 can be considered as low quality; sample RP-4 as acceptable; samples RP-6 and RP-8 as good. In the samples analyzed, the most abundant flavonoids were chrysin, pinocembrin, galangin and pinobanksin-3-O-acetate.
Regarding phenolic acids, caffeic acid, p-coumaric acid and ferulic acid were the most abundant ones. As for phenolic acid derivatives, 3,4-dimethyl caffeic acid (DMCA), caffeic acid prenyl, benzyl, phenylethyl (CAPE) and cinnamyl esters were the most representative compounds. 4. Conclusions An efficient closed-vessel MAE process was developed and optimized for the first time by means of a response surface experimental design methodology for the fast extraction of phenolic acids and flavonoids from raw propolis. Compared with traditional methods, the proposed MAE procedure allowed a reduction of extraction time and solvent use. The extraction efficiency of MAE was comparable with that obtained with conventional extraction methods and UAE, without degradation of the compounds of interest.
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The results of the HPLC analysis of real matrixes, carried out using a validated method based on the use of a fused-core C18 column, indicated a great variability in the content of the active compounds. In this context, the extraction procedure and HPLC method developed in this study can be considered reliable and useful tools for the comprehensive multi-component analysis of polyphenols in extracts of raw propolis to be used in apitherapy. Acknowledgments The authors are grateful to Kontak (Pozzo d’Adda, Milan, Italy) for the financial support. The authors acknowledge the “Fondazione Cassa di Risparmio di Modena” for funding the HPLC–ESI–MS system at the Centro Interdipartimentale Grandi Strumenti (CIGS) of the University of Modena and Reggio Emilia. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.jpba.2013.04.003. References [1] J.M. Sforcin, V. Bankova, Propolis: is there a potential for the development of new drugs? J. Ethnopharmacol. 133 (2011) 253–260. [2] A.M. Gómez-Caravaca, M. Gómez-Romero, D. Arráez-Román, A. SeguraCarretero, A. Fernández-Gutiérrez, Advances in the analysis of phenolic compounds in products derived from bees, J. Pharm. Biomed. Anal. 41 (2006) 1220–1234. [3] Y. Xu, L. Luo, B. Chen, Y. Fu, Recent developments of chemical components in propolis, Front. Biol. China 4 (2009) 385–391.
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