Anal. Chem. 2007, 79, 6767-6774
Simultaneous Ultrasound-Assisted Emulsification-Extraction of Polar and Nonpolar Compounds from Solid Plant Samples J. A. Pe´rez-Serradilla, F. Priego-Capote, and M. D. Luque de Castro*
Department of Analytical Chemistry, Annex Marie Curie Building, Campus of Rabanales, University of Co´ rdoba, E-14071 Co´ rdoba, Spain
A new approach to solid sample preparation for the simultaneous isolation of polar and nonpolar compounds using a microemulsion as leaching medium is proposed. Methanol/water (dispersed-phase)-hexane (continuousphase) emulsions formed in the presence of ultrasound and a solid sample allow polar and nonpolar compounds to be transferred to the dispersed and continuous phase, respectively. The efficiency of this dual sample preparation approach was assessed in the characterization of natural products of variable hardness including acorns, grape seeds, and alperujo (a residue of olive oil production). The time needed for quantitative extraction of the target fractions (phenol compounds and fatty acids) is 9 min for acorns and alperujo and 20 min for grape seeds; the longer time needed for grape seeds can be attributed to higher matrix hardness. Such good performance can be ascribed to the ultrasound-enhanced formation of methanol/water microdroplets 1-15 µm in size, which act as solid-liquid microextractors spanning a large surface area. The presence of the sample was found to greatly improve emulsion stability, which can be ascribed to the amphiphilic nature of the fatty acids in the samples. Following leaching and separation of the two phases by centrifugation, the polar and nonpolar fractions were analyzed by HPLC-diode array detection and GC/MS, respectively. The proposed approach provides extraction efficiency similar to the Folch method (reference method for fat extraction, 4.5 h) in a shorter time and extraction efficiency equal to or higher than the stirring-based method (reference method for phenol compounds extraction, 24 h). Optimizing sample preparation (particularly with solid samples) remains a priority goal and a subject of growing importance in analytical chemistry1,2 as redoubled efforts are being made to bridge the performance gap with other steps of the analytical process such as detection or data processing. * Corresponding author. Tel. and Fax: +34 957 218615. E-mail:
[email protected]. (1) Mester, Z.; Sturgeon, R. Comprehensive Analytical Chemistry; Elsevier: Amsterdam, 2003. (2) Luque de Castro, M. D.; Priego-Capote, F. Analytical Applications of Ultrasound; Elsevier: Amsterdam, 2006. 10.1021/ac0708801 CCC: $37.00 Published on Web 07/10/2007
© 2007 American Chemical Society
The use of ultrasonic energy in various steps of the analytical process (e.g., leaching, liquid-liquid extraction, slurry formation, emulsification, filtration) provides widely documented advantages.2 Ultrasound-assisted emulsification was first reported by Wood and Loomis in 1927.3 The most widely accepted mechanism for ultrasound-assisted emulsification is based on the effects of cavitation, which are deemed crucial for the process to develop. The implosion of bubbles generated by cavitation causes intensive shock waves in the surrounding liquid and formation of liquid jets of a high velocity, which can cause droplet disruption in the vicinity of collapsing bubbles.4-6 Those factors with a favorable effect on cavitation in liquids generally improve emulsification by reducing droplet size in the dispersed phase right after disruption. The beneficial effects of ultrasonic assistance to emulsification and the underlying mechanisms have been the subject of extensive research and aroused increasing interest in the process.7 In fact, a number of methods where ultrasonic emulsification is a key step in the determination of the target analytes have been proposed. Most such methods are focused on the determination of metallic elements in liquid organic samples using atomic detectors.8,9 Ultrasound is also known to facilitate extraction processes.2 The effects of ultrasound on leaching are also essentially primarily related to cavitation. Thus, the implosion of bubbles formed during ultrasound application produces rapid adiabatic compression of gases and vapors within the bubbles or cavities and a highly efficient temperature and pressure as a result. The increased temperature enhances the solubility of the analytes in the leachant and facilitates their diffusion from the sample matrix to the outer region; on the other hand, the increased pressure facilitates penetration of the leachant into the sample matrix and transfer from the matrix to liquid phase at the interface.10 Various types of analytes including polar11,12 and nonpolar13,14 organic com(3) Wood, R. W.; Loomis, A. L. Phil. Mag. 1927, 4, 417-436. (4) Lauterborn, W.; Ohl, C. D. Ultrason. Sonochem. 1997, 4, 65-75. (5) Li, M. K.; Fogler, H. S. J. Fluid. Mech. 1978, 88, 499-511. (6) Li, M. K.; Fogler, H. S. J. Fluid. Mech. 1978, 88, 513-528. (7) Behrend, O.; Ax, K.; Schubert, H. Ultrason. Sonochem. 2000, 7, 77-85. (8) Murillo, M.; Gonzalez, A.; Ramı´rez, A.; Guille´n, N. At. Spectrosc. 1994, 15, 90-95. (9) Wang, T. B.; Jia, X. J.; Wu, J. J. Pharm. Biomed. Anal. 2003, 33, 639-646. (10) Luque-Garcı´a, J. L.; Luque de Castro, M. D. Trends Anal. Chem. 2003, 22, 41-47. (11) Palma, M.; Barroso, C. G. Anal Chim. Acta. 2002, 458, 119-130. (12) Alonso-Salces, R. M.; Barranco, A.; Corta, E.; Berruela, L. A.; Gallo, B.; Vicente, F. Talanta 2005, 65, 654-662.
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pounds, organometals,15,16 and metallic elements17,18 have thus been efficiently leached from solid samples. Ultrasound equipment for leaching usually consists of a commercial ultrasonic bath or probe. Comparative studies have revealed that probes afford more expeditious leaching than ultrasonic baths do.19 These two applications of ultrasonic energy (namely, emulsification and extraction) can be combined into a single approach for the simultaneous extraction of both polar and nonpolar compounds from a solid sample by bringing them into close contact with two extractants of disparate polarity. The small droplets in the microemulsion can act as microextractors with a high mass-transfer surface, thus expediting the leaching kinetics of the dispersed phase. Following mass transfer, the two phases can be readily separated for subsequent analysis by salting out or centrifugationsthe latter is to be preferred as it avoids the presence of foreign substances in the extract. For the extraction of the fatty fraction from solid samples, the AOAC recommends reflux-extraction procedures such as Goldfisch and Soxhlet, but also the Folch method20 is frequently used.21 Quantitative removal is achieved by these methods, but they require long process times that can range from 8 to 24 h. The extractant most frequently used has been n-hexane,22-24 although petroleum ether25 and hexane-2-propanol26 have also been used. Extraction of the phenolic fraction from solid samples is usually carried out by stirring27-29 using methanol-water mixtures.30,31 Auxiliary energies such as ultrasound32,33 and microwaves,21,34 and superheated liquids35,36 and supercritical fluids (mainly CO2)37,38 as extraction techniques have been used to accelerate the removal (13) Chu, S. G.; Hong, C. S.; Rattner, B. A.; McGowan, P. C. Anal. Chem. 2003, 75, 1058-1066. (14) Sulkowsky, W.; Rosinska, A. J. Chromatogr., A 1999, 845, 349-355. (15) Tu, Q.; Qian, J.; Frech, W. J. Anal. At. Spectrom. 2000, 15, 1583-1588. (16) Nischwitz, V.; Michalke, B.; Kettrup, A. Analyst 2003, 128, 109-115. (17) Liva, M.; Olivas, R. M.; Ca´mara, C. Talanta 2000, 51, 381-387. (18) Lo´pez-Garcı´a, I.; Campillo, N.; Arnau-Jerez, I.; Herna´ndez-Co´rdoba, M. Anal. Chim. Acta 2005, 531, 125-129. (19) Aleixo, P. C.; Ju´nior, D. S.; Tomazelli, A. C.; Rufino, I. A.; Berndt, H.; Drug, F. J. Anal. Chim. Acta 2004, 512, 329-337. (20) Folch, J.; Lees, M.; Sloane-Stanley, G. H. Biol. Chem. 1957, 226, 497-509. (21) Perez-Serradilla, J. A.; Ortiz, M. C.; Sarabia, L.; Luque de Castro, M. D. Anal. Bioanal. Chem. 2007, 388, 451-462. (22) Elkhori, S.; Pare, J. R. J.; Belanger, J. M. R.; Perez, E. J. Food Eng. 2006, 79, 1110-1114. (23) Nebel, B. A.; Mittelbach, M. Eur. J. Lipid Sci. Technol. 2006, 108, 398403. (24) Rao, Y. R.; Jena, K. S.; Sahoo, D.; Rout, P. K.; Ali, S. J. Am. Oil Chem. Soc. 2005, 82, 749-752. (25) Kays, S. E.; Archibald, D. D.; Sohn, M. J. Sci. Food Agric. 2005, 85, 15961602. (26) Gunnlaugsdottir, H.; Ackman, R. G. J. Sci. Food Agric. 1993, 61, 235-40. (27) Spigno, G.; Tramelli, L.; De Faveri, D. M. J. Food Eng. 2007, 81, 200-208. (28) Luthria, D. L.; Mukhopadhyay, S.; Kwansa, A. L. J. Sci. Food Agric. 2006, 86, 1350-1358. (29) Nepote, V.; Grosso, N. R.; Guzman, C. A. J. Sci. Food Agric. 2005, 85, 3338. (30) Japo´n-Luja´n, R.; Luque-Rodrı´guez, J. M.; Luque de Castro, M. D. Anal. Bioanal. Chem. 2006, 385, 753-759. (31) Turkmen, N.; Velioglu, Y. S. J. Sci. Food Agric. 2007, 87, 1408-1416. (32) Japo´n-Luja´n, R.; Luque-Rodrı´guez, J. M.; Luque de Castro, M. D. J. Chromatogr., A 2006, 1108, 76-82. (33) Ruiz-Jimenez, J.; Priego-Capote, F.; Luque de Castro, M. D. J. Chromatogr., A 2004, 1045, 203-210. (34) Liazid, A.; Palma, M.; Brigui, J.; Barroso, C. G. J. Chromatogr., A 2007, 1140, 29-34. (35) Luque-Rodrı´guez, J. M.; Luque de Castro, M. D.; Pe´rez-Juan, P. Talanta 2005, 68, 126-130. (36) Japo´n-Luja´n, R.; Luque de Castro, M. D. J. Agric. Food Chem. 2007, 55, 3629-3634.
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of the polar or nonpolar fraction from solid samples, although, as far as the authors know, a simultaneous leaching of the two fractions has not previously been reported. The aim of this work was to develop an ultrasound-assisted method for the simultaneous leaching of polar and nonpolar compounds from solid samples by using an emulsion of n-hexane and a methanol-water mixture for the quantitative removal of the two fractions in as short a time as possible. The solid samples studied included acorns (the main ingredient of the diet of Iberian pigs),39 alperujo (a residue from the two-phase decantation process used by the olive oil industry),40 and grape seeds (which account for ∼15% of the solid waste produced by wine industries.35 The analytes in the nonpolar fraction (fatty acids) were determined by GC/MS after derivatization and those in the polar fraction (phenol compounds) by HPLC-diode array detection (DAD). The Folch method and the stirring extraction method were also used to extract the nonpolar and polar compounds, respectively, from the samples under study, and the results thus obtained were compared with those provided by the proposed method in terms of extract composition, leaching efficiency, and extraction time. EXPERIMENTAL SECTION Instruments and Apparatus. Ultrasonic irradiation was applied with a Branson 450 digital sonifier (20 kHz, 400 W) equipped with a cylindrical titanium alloy probe (12.70-mm diameter), which was directly immersed in the emulsificationextraction beaker. A 1250 rpm magnetic stirrer (Berghof, Germany) was also used to ensure contact between the solid and emulsion. A centrifuge (Selecta, Barcelona, Spain) was used to break the emulsions. A DM5000B optical microscope from Leica Microsystems (Wetzlar, Germany) coupled with a DC500 Leica digital camera was used to measure droplet size in the emulsion. A vortex from Ika-Works (Wilmington, NC) and the centrifuge were used to derivatize fatty acids into their methyl esters. Fatty acids were determined on a Varian CP 3800 gas chromatograph coupled to a Saturn 2200 ion trap mass spectrometer (Sugar Land, TX) furnished with a VF-23ms FactorFour capillary column (60 m × 0.25 mm, 0.25 µm), also from Varian. This column affords the separation of the cis and trans isomers of FAMEs, if present, in the extracts. An Agilent 1100 liquid chromatograph consisting of a G1322A vacuum degasser, a G1315A DAD, and a Rheodyne 7725 highpressure manual injection valve (20-µL injection loop) was used to analyze the phenolic fraction by HPLC. The analytical columns used were a Lichrospher 100 RP-18 (250 mm × 4 mm i.d. 5 µm) and an Ultrabase C-18 (250 mm × 4.6 mm i.d., 5 µm), both purchased from Ana´lisis Vı´nicos (Ciudad Real, Spain). A Kromasil 5 C-18 precolumn (15 mm × 4.6 mm i.d., 5 µm) protected with a steel holder, both from Scharlab, (Barcelona, Spain), was used to clean up extracts. (37) Sparks, D.; Herna´ndez, R.; Zappi, M.; Blackwell, D.; Fleming, T. J. Am. Oil Chem. Soc. 2006, 83, 885-891. (38) Chen, J. L.; Liu, C. Y. Anal. Chim. Acta 2005, 528, 83-88. (39) Lo´pez-Bote, C. J. Meat Sci. 1998, 49, 17-27. (40) Priego-Capote, F.; Ruiz-Jime´nez, J.; Luque de Castro, M. D. J. Chromatogr., A 2003, 1045, 239-246.
Operational variables were optimized by using the software Statgraphics plus v.2.1 for Windows (Stat Point, Inc., Herndon, VI). Chemicals. All reagents were analytical grade or higher. Methanol, n-hexane, trichloromethane, formic acid, and acetonitrile were provided by Scharlab, and acetic acid was from Panreac (Barcelona, Spain). A 2:1 (v/v) trichloromethane-methanol mixture was used in the Folch extraction; NaCl and NaClO4 were used as salting-out agents to facilitate phase separation and anhydrous Na2SO4 (Panreac) was employed as a drying agent of the nonpolar phase. Sodium methylate (0.5 M) in methanol was used as the derivatizating reagent to obtain FAMEs. All applicable safety precautions (viz. gloves, mask, fume hood, an isolated sound-proof cabinet for ultrasonication) were adopted. Deionized water (18 MΩ‚cm) from a Millipore Milli-Q water purification system was used to prepare both the 80:20 (v/v) methanol-water extractant mixture and the mobile phases for liquid chromatography. A multistandard containing the methyl esters of 19 fatty acids (viz. lauric acid (12:0), tridecanoic acid (13:0), myristic acid (14: 0), pentadecanoic acid (15:0), palmitic acid (16:0), trans-palmitoleic acid (t16:1), palmitoleic acid (16:1), heptadecanoic acid (17:0), stearic acid (18:0), elaidic acid (t18:1), oleic acid (18:1), trans,trans-linoleic acid (tt18:2), cis,trans-linoleic acid (ct18:2), trans,cis-linoleic acid (tc18:2), linoleic acid (18:2), linolenic acid (18:3), arachidic acid (20:0), cis-11-eicosenoic acid (20:1), and behenic acid (22:0)), all from Sigma-Aldrich (St. Louis, MO), was used. Undecanoic acid methyl ester from Fluka (Steinheim, Germany) was used as external standard in GC/MS determinations. Gallic acid (Merck, Darmstadt, Germany); catechin, epicatechin, procyanidin B2, and ellagic acid (Sigma, Steinheim, Germany); and tyrosol, hydroxytyrosol, and verbascoside (Extrasynthese, Genay, France) were used as standards in the HPLCDAD determinations. Samples. Quercus ilex acorns were collected in Andalucı´a, Spain; both grape seeds and alperujo samples were obtained from a winery and olive oil industry in Montilla (Co´rdoba, Spain). All samples were dried, milled, sieved to a 0.5-mm particle size, and stored at 4 °C until use. The studied samples were intended to be representative of variable hardness and density. Ultrasound-Assisted Emulsification-Extraction (UAEE). Two grams of sample (acorns, alperujo, or grape seeds), 30 mL of n-hexane, and 25 mL of 80:20 (v/v) methanol-water mixture were placed in a 100-mL beaker (46-mm diameter), which was then immersed in a water bath at room temperature to facilitate heat dissipation. The probe was directly dipped into the solidliquid system, which was magnetically stirred. The experimental setup is depicted in Figure 1. Ultrasonic irradiation (irradiation duty cycle 0.8 s/s, output amplitude 10% of the converter, applied power 450 W with the probe placed at 0.5 cm from the top surface of the liquid phase) was applied for a preset time (9 min for both acorns and alperujo, and 20 min for grape seeds) in order to facilitate the emulsion formation and leaching of the target compounds to the appropriate phase as a function of their polarity. After leaching, emulsions were disrupted by centrifugation at 4000 rpm (centrifugal force 2100g) for 5 min. The aqueousmethanol phase and n-hexane phase were separatedsand the
Figure 1. Experimental setup for simultaneous ultrasound-assisted emulsification-extraction.
organic solvent was evaporated under an N2 streamsfor subsequent HPLC-DAD and GC/MS analysis, respectively. Reference Method for Fat Extraction. Folch extraction20 was selected as the reference method for fat extraction by virtue of its mild working conditions, which avoid potential alterations of extracted fat. An amount of 25 g of sample was put into contact with 75 mL of 2:1 (v/v) trichloromethane-methanol and subjected to magnetic stirring in a 250-mL Erlenmeyer flask for 45 min. Then, the mixture was filtered in a Buchner device and the solid phase re-extracted three more times with the same volume of extractant. The liquid phases were combined in a separatory funnel and supplied with 35 mL of saturated sodium chloride in water and 0.5 g of NaClO4, and the mixture was shaken gently. After phase separation, the chloroform phase was dried with sodium sulfate and filtered. Finally, the extractant was evaporated under an N2 stream. The total time required was 270 min. Reference Method for Extracting Phenolic Compounds.32 An amount of 1 g of dried, milled sample and 12.5 mL of 80:20 (v/v) methanol-water mixture was placed in a beaker and stirred at 40 °C for 24 h, after which the residue was removed by centrifugation prior to analysis by liquid chromatography. Preparation of Fatty Acid Methyl Esters. An amount of 0.05 g of fat extract was diluted to 5 mL with n-hexane and homogeneized in a vortex for 30 s; then, 0.25 mL of sodium methylate in methanol was added and the mixture shaken vigorously in a vortex for 3 min and centrifuged at 4000 rpm (centrifugal force 2100g) for 2 min. The resulting supernatant was transferred to a test tube and evaporated to dryness under an N2 stream. A volume of 2 mL of n-hexane was used to reconstitute the residue, which was shaken in a vortex for 1 min. Following 1:10 dilution, a 1-µL aliquot of the solution thus obtained was supplied with the external standard and injected into the GC/MS system. GC/MS Separation and Detection. This step was optimized by using the FAMEs multistandard. Methyl undecanoate was used as external standard (150 µL of 1000 µg g-1 solution in n-hexane was added prior to analysis) by virtue of its physical and chemical Analytical Chemistry, Vol. 79, No. 17, September 1, 2007
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Figure 2. GC chromatogram for the nonpolar fraction of acorns as obtained using the UAEE method under optimal working conditions. The magnified detail of the chromatogram exposes the peaks of the less abundant analytes.
properties being similar to those of the derivatized analytes and its absence in the samples. Helium at a constant flow rate of 1 mL min-1 was used as carrier gas for the GC/MS determination of FAMEs. The temperature program of the column was as follows: 50 °C (hold 2 min) and 5 °C min-1 ramp to 250 °C (hold 15 min). Injections (1 µL each) were done in the split mode (ratio of 1), using an injector temperature of 250 °C. The ion trap mass spectrometer was operated in the electron impact ionization (EI) positive mode. For EI measurements, the instrumental parameters were set as follows: filament emission current 80 µA, electron multiplier voltage 1600 V, modulation amplitude 4 V, perfluorotributylamine (FC-43) as reference, and multiplier offset 200 V. The transfer line and ion trap were set at 170 °C and the manifold at 50 °C. The storage window was set between 40 and 600 m/z, and the selected-ion monitoring ion preparation mode was used. The scan time for data acquisition was set at 1.0 s, with 3 microscans/s. Complete separation of FAMEs was accomplished within 40 min (see Figure 2). The analytes were identified by comparison with both retention times and mass spectra for the multistandard. Calibration graphs plots in the form of peak area versus standard concentration of each compound were run for all analytes. Limits of detection (LODs) and quantification (LOQs)sexpressed as the mass of analyte that gives a signal that is 3σ and 10σ, respectively, above the mean blank signal, where σ is the standard deviation of the blank signal for 10 measurementssranged from 0.08 to 0.76 µg g-1 and from 0.26 to 2.51 µg g-1, respectively. The linear dynamic ranges for all compounds were between the LOQ and 10.000 µg g-1. The correlation coefficients of the regression lines ranged from 0.9979 to 0.9999. HPLC-DAD Separation-Detection. The polar extracts were injected into the chromatograph after microfiltration. No 6770
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preconcentration was required. Acorns and alperujo polar phases were analyzed using the Lichrospher HPLC column, and the grape seed polar phase with the Ultrabase HPLC column. The eluents and gradient programs used are shown in Figure 3. A flow rate of 1 mL min-1 was employed in all cases. All chromatograms were acquired at 280 nm and obtained within 50 min. The polar fractions from the three samples were analyzed for various phenols. Thus, gallic and ellagic acids were identified in that from acorns; hydroxytyrosol, tyrosol, and verbascoside in that from alperujo; and gallic acid, catechin, procyanidin B2, and epicatechin in the grape seed polar extracts. The LODs for these phenol compounds ranged from 1.4 to 4.3 µg g-1 and their LOQs from 3.4 to 14.3 µg g-1. Correlation coefficients for the regression lines ranged from 0.9970 to 0.9999. RESULTS AND DISCUSSION Preliminary Studies. Mixing two immiscible liquids to obtain an emulsion almost invariably requires the assistance of energy; also, the resulting emulsion is usually unstable. Stability is especially low for liquid-liquid dispersions with a high interfacial area. In this respect, the use of additives such as emulsifiers allow finely dispersed media to be easily created. The most common class of emulsifiers is that of surfactants, which by virtue of their amphiphilic structure are easily adsorbed at the interface between two phases, thereby helping stabilize the droplets of the dispersed phase of an emulsion. We used n-hexane22-24 and 80:20 (v/v) methanol-water,30,31 which have proved excellent extractants for nonpolar and polar compounds, respectively. A univariate optimization study was conducted in order to identify the most suitable 80:20 (v/v) methanol-water/n-hexane ratio to form an emulsion in which the diameter of the dispersed-phase droplets was as low as possible
Figure 3. HPLC-DAD chromatograms for the polar fraction of acorns, alperujo, and grape seeds as obtained using the UAEE method under optimal working conditions and recorded at 280 nm. The eluent and gradient program used in each chromatographic separation are also shown. HAc, acetic acid; NaAc, sodium acetate; ACN, acetonitrile.
for a more stable emulsion. Test were carried out in the presence of acorns; medium values of probe position, ultrasound radiation amplitude, and percent of ultrasound exposure duty cycleswhich have a slight influence on the emulsification processswere selected. A 5:6 (v/v) ratio, with which the emulsion formed completely within the first 10 s of ultrasound application, was selected as optimal and adopted for subsequent tests. The range tested in this univariate study was from 4:6 to 6:4 (v/v). The presence of the sample was found to greatly improve stability in the emulsions. This can be ascribed to the amphiphilic nature of the fatty acids in the samples (with a hydrophilic carboxylic part and a hydrophobic long organic chain), which
facilitates adsorption at the surface of the polar microdroplets via the carboxylic group, the organic chain remaining in the n-hexane continuous phase. Thus, fatty acids act as emulsifiers and facilitate formation and stabilization of the emulsions. Restrictive working conditions referred to sample amount and sample/extractant volume ratio are as follows: (1) sample amount: as a function of the stirrer power in order to maintain the solid in suspension; (2) volume of liquid phase enough to maintain the sample particles in suspension and avoid the contact between the ultrasound probe and the magnetic bar. On the other hand, in order to ensure a whole emulsion formation (for which the presence of sample is needed), the volume of liquid phase Analytical Chemistry, Vol. 79, No. 17, September 1, 2007
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Figure 4. Droplets formed during emulsification as seen under a light microscope. Each division of the scale corresponds to 10 µm.
must not exceed the level to which the stirrer disperses the particles. As can be seen in Figure 4 for acorns, the diameters of the methanol-water droplets (dispersed phase) formed in the emulsification process spanned the range 1-15 µm. The characteristics of the emulsions obtained after ultrasound application in the presence of alperujo or grape seeds were similar to those of the emulsions obtained with acorns. Note that a stable emulsion was formed from the very beginning of the extraction process; therefore, if one assumes the extracted fat acts as a stabilizer, a small amount of fat is enough to stabilize the emulsion. Optimization of the Ultrasound-Assisted EmulsificationExtraction Process. Using the extractants ratio selected in the preliminary studies in the presence of sample, the emulsion is formed in few seconds after ultrasound application, so the emulsion exists since the beginning of the extraction. The influence of the main factors involved in the leaching process was studied by using an experimental design that was applied to each type of sample. Two response variables were examined in each case, namely, the total amount of fatty acids and that of phenol compounds (those identified in each sample). The optimization study involved a complete two-level factorial design allowing 5 degrees of freedom and involving 16 randomized runs plus 3 central points; the design was used to screen the 4 factors with a potential influence on the extraction process,2 namely: radiation amplitude (tested range 10-30%), duty cycle (30-80%), irradiation time (2-7 min), and probe depth (0.5-1.5 cm from the surface of the solid-liquid system). The results obtained in the screening studies for the three types of sample were similar, which testifies to the robustness of the proposed method. The main conclusion was that the sole factor significantly affecting the extraction efficiency of polar and nonpolar compounds was the irradiation time, as it has a positive and significant influencesat 95% confidence level, its influence is out of the experimental errorsin the two response variables. As regards the effects of other factors, the extraction efficiency was higher with the highest ultrasound exposure duty cycle (80%)s positive influence, although not significant at 95% confidence levelsand the minimum probe depth and radiation amplitude tested (0.5 cm and 10%, respectively)snegative influence, although not significant at 95% confidence level. A univariate study of the influence of the extraction time on the extraction efficiency of polar and nonpolar compounds was 6772 Analytical Chemistry, Vol. 79, No. 17, September 1, 2007
Figure 5. Kinetics of extraction of fatty acids (-b-) and phenol compounds (- - -). The extraction efficiency is expressed as a percentage of total fatty acids or total phenolic compounds extracted with the proposed method; the maximum amount extracted in each kinetic curve was taken to be 100%.
carried out on each sample in order to obtain a better understanding of the extraction kinetics. Figure 5 shows the variation of the extraction efficiency for fatty acids and phenol compounds with the time for the three types of samples studied. The maximum extraction efficiency for fatty acids and phenols in acorns and alperujo was obtained with 7 and 9 min, respectively; the times needed to obtain similar results with grape seeds were 15 and 20 min, respectively. Therefore, the optimal working conditions were those described in the Experimental Section. The longer time required for quantitative extraction of phenols and fatty acids from grape seeds as compared to the extraction time in the case of acorns and alperujo can be attributed to its high matrix hardness, which is more resistant to ultrasonic energy. The variable influence of humidity of the different samples was avoided by previous sample drying. The phenol compounds extracted in the kinetics tests from the different samples exhibited a similar behavior; thus, their concentrations in the extracts decreased after maximal extraction. This can be ascribed to the high temperatures and pressures created by the effect of microbubble implosion during cavitation. In fact, when ultrasound energy is applied to a liquid medium,
Table 1. Percentage of Fatty Acids in the Nonpolar Fraction from Acorns, Alperujo, and Grape Seeds As Obtained Using the UAEE and Folch Extraction Methodsa acorns
alperujo
grape seeds
fatty acid
proposed method
Folch method
proposed method
Folch method
proposed method
Folch method
14:0 15:0 16:0 16:1 17:0 18:0 18:1 18:2 18:3 20:0 20:1 22:0 total fat
0.11 (0.50) 0.08 (1.22) 17.90 (0.24) 0.20 (1.89) 0.27 (0.76) 3.25 (1.23) 55.40 (3.44) 20.88 (0.94) 1.29 (0.22) 0.26 (1.02) 0.29 (1.63) 0.08 (0.02) 9.59 (0.57)
0.13 (1.48) 0.11 (1.79) 15.95 (0.66) 0.37 (0.84) 0.7 (0.05) 5.35 (2.21) 57.17 (0.55) 18.49 (1.85) 0.98 (0.24) 0.25 (1.88) 0.44 (2.01) 0.06 (1.52) 9.54 (0.44)
ndb nd 8.26 (1.15) 0.54 (1.64) 0.09 (0.54) 1.85 (2.54) 79.18 (0.94) 9.34 (1.86) 0.42 (3.95) 0,16 (2.98) 0.16 (1.64) nd 20.20 (2.11)
nd nd 8.20 (2.94) 0.45 (1.14) 0.09 (1.47) 1.69 (0.87) 79.33 (1.23) 9.45 (2.33) 0.42 (2.9) 0.18 (0.54) 0.18 (0.20) nd 20.50 (1.25)
nd nd 7.46 (1.75) 0.17 (0.46) 0.11 (3.91) 3.52 (1.31) 26.44 (0.63) 61.64 (3.39) 0.36 (0.19) 0.13 (2.65) 0.17 (1.94) nd 7.91 (0.88)
nd nd 7.22 (2.45) 0.2 (0.80) 0.09 (1.91) 3.49 (2.78) 26.29 (1.94) 62.02 (2.54) 0.39 (0.99) 0.095 (3.10) 0.22 (1.25) nd 7.94 (1.34)
a The percentage of total fat extracted from each type of sample is also shown. Errors, in parentheses, are expressed as percent relative standard deviations (n ) 3 replicates). b nd, not detected.
cavitation produces high pressures and high localized temperatures, which has a favorable effect on the solid-liquid extraction kinetics; once the maximum extraction efficiency is reached, excess ultrasound energy causes sonolysis of the solvent.2 Since most phenol compounds present in natural samples have antioxidant properties,41 the decrease in concentration observed after the maximum extraction efficiency was reached can be ascribed to a degradation process in which free radicals generated by sonolysis of the extractant react with extracted phenol compounds. This entails performing an exploratory kinetic study before the proposed methodology is applied. Comparison of the Fractions Obtained with the Proposed Method and the Extracts Provided by the Reference Methods. The total fat and phenol contents obtained with the Folch and phenol reference extraction methods, respectively, were compared with those provided by the UAEE method under optimal working conditions (Tables 1 and 2). Based on the results of Table 1, the Folch method and the proposed method provided similar fat contents for all samples. Also, the leaching efficiency for phenol compounds obtained with the proposed method was equal to or higher than that provided by the reference extraction method (Table 2). The composition of the nonpolar fractions obtained from the three types of samples studied under the optimal working conditions for the proposed method were compared with those provided by the reference Folch method; by way of example, Figure 2 shows the chromatogram for the nonpolar fraction from the acorns. The relative concentration of each fatty acid in the nonpolar extractsstaking as 100% the overall concentration of fatty acids in each extractsis shown in Table 1. Similarly to Folch extracts, no trans fatty acids were detected in the nonpolar fractions obtained by applying the proposed method to acorns, alperujo, or grape seeds. Taking into account the mild conditions used in the Folch method, no cis-trans transformation during extraction is to be expected from it or from the proposed method either. (41) Santos-Buelga, C.; Williamson, G. Methods in Polyphenol Analysis; The Royal Society of Chemistry: Cambridge, UK, 2003.
Table 2. Contents of the Target Phenols in Each Type of Samplea phenolic compound
proposed method
REMPb
gallic acid ellagic acid
Acorns 105.7 (3.89) 168.2 (4.07)
106.1 (3.12) 167.4 (3.58)
hydroxytyrosol tyrosol verbascoside
Alperujo 366.6 (2.56) 621.0 (3.33) 52.1 (2.13)
202.1 (2.51) 389.0 (2.96) 47.9 (2.40)
gallic acid catechin procyanidin B2 epicatechin
Grape Seeds 18.2 (2.46) 35.4 (1.88) 26.2 (3.25) 35.7 (2.83)
17.1 (2.43) 37.8 (2.67) 15.5 (2.76) 34.6 (2.24)
a Expressed in µg g-1. Errors, in parentheses, are expressed as percent relative standard deviations (n ) 3 replicates). b REMP, reference extraction method for phenols.
A paired-sample comparison t-test was used to determine whether both methods provided similar results at the 95% confidence level. The null hypothesis was that both methods gave identical results or, in other words, that the differences between the Folch and UAEE results were not significant. The calculated t-values were smaller than the theoretical value at R ) 0.05 and 12 degrees of freedom for all the samples, so the null hypothesis was accepted. This means that, at the selected significance level, the differences between the composition of the nonpolar fractions obtained by the two methods were within the experimental error. Precision of the Proposed Method. The precision of the proposed method in terms of within-laboratory reproducibility and repeatability was assessed by using a single experimental setup with duplicates.42 Tests involved extracting 2 g of sample under the optimal working conditions. Two measurements of nonpolar and polar extracts per sample and day were performed on 7 days. The repeatability and within-laboratory reproducibility for fatty (42) Massart, D. L.; Vanderginste, B. G. M.; Buydens, L. M. C.; De Jong, S.; Lewi, P. J.; Smeyers-Verbeke, J. Handbook of Chemometrics and Qualimetrics, Part A; Elsevier: Amsterdam, 1997.
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acids in the nonpolar fractions, both expressed as relative standard deviation, were ranged from 3.78 to 5.76% and from 7.32 to 10.54%, respectively. For the phenol compounds in the polar fractions, the repeatability and within-laboratory reproducibility were 3.018.57 and 7.05-17.34%, respectively. It should be noted that the precision for grape seeds was slightly poorer than those for acorns and, especially, alperujo. This can be ascribed to the ease of solvent penetration in the samples decreasing with increasing hardness. CONCLUSIONS Ultrasound-assisted emulsification-extraction has for the first time been used for the simultaneous removal of polar and nonpolar compounds from solid samples. The proposed method allows the target analytes to be leached in a very short time (9-20 min, depending on the sample) as compared with the reference extraction method for phenol compounds (24 h) and the Folch method for fat isolation (4.5 h). The total fat content and the composition of the nonpolar extracts from the three types of samples studied were similar to those obtained with the Folch method. Also, the leaching efficiency of the proposed method for phenolic compounds was equal to or higher than that of the reference extraction method. The performance of the proposed approach can be ascribed mainly to the following factors: (1) The high contact area between
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the solid sample and the dispersed and continuous phases as a consequence of ultrasound-assisted emulsification; and the high surface area of the small polar droplets, which act as microextraction units and facilitate the leaching of phenol compounds with a high efficiency. (2) Concerning the continuous nonpolar phase, the fact that fatty acids migrate to the interface by virtue of their amphiphilic nature leads to a lower energy state, which stabilizes the emulsion and a displacement of the solid-hexane leaching equilibrium of fatty acids. (3) The high effective localized temperature and pressure generated by cavitation, which expedites the solid-liquid extraction kinetics. ACKNOWLEDGMENT Spain’s Ministry of Science and Technology is gratefully acknowledged for financial support (Project CTQ2006-01614). J.A.P.-S. and F.P.-C. are also grateful to Spain’s Ministry of Education for award of their research training fellowships. P. Go´mez and D. Gracia, of the Microscopy Unit (SCAI) of the University of Cordoba, are gratefully acknowledged for their cooperation.
Received for June 11, 2007. AC0708801
review
April
30,
2007.
Accepted
