tandem mass spectrometry studies of the phenolic compounds in honey

Journal of Chromatography A, 1216 (2009) 6620–6626

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Liquid chromatography/tandem mass spectrometry studies of the phenolic compounds in honey夽 Magdalena Biesaga ∗ , Krystyna Pyrzynska Department of Chemistry, University of Warsaw, Pasteura 1, 02-093 Warsaw, Poland

a r t i c l e

i n f o

Article history: Received 18 March 2009 Received in revised form 21 July 2009 Accepted 30 July 2009 Available online 3 August 2009 Keywords: Honey Flavonoids Phenolic acids Mass spectrometry

a b s t r a c t An improved and simplified analytical method which offers rapid, accurate determination and identification of phenolic compounds in honey samples is reported. The honey samples were diluted by acidified water (pH 2), and analyzed by HPLC–ESI-MS/MS. Simultaneously determination of phenolic acids and flavonoids was carried out by a liquid chromatography–mass spectrometry. Comparison between atmospheric pressure chemical ionization (APCI) and electrospray ionization (ESI) was performed by analysis of standards. Fragmentation of analytes for subsequent selective Multiple Reaction Monitoring (MRM) analysis was investigated in negative mode. Sample preparation without separation of sugars and clean-up procedure, followed by fast chromatographic separation using a narrow-bore column C18 (50 mm × 2.1 mm, 3 ␮m) allowed the analysis to be completed in a short run time. LODs were ranged between 1 and 15 ng L−1 for p-coumaric acid and naringenin, respectively. The method was applied for determination of phenolic acids and flavonoids in honey samples from different botanical origin. © 2009 Elsevier B.V. All rights reserved.

1. Introduction There is a lack of knowledge about the profiles of phenolic substances in honey from various sources in order to evaluate the quality of the product. Several studies have been demonstrated that honey serves as a source of natural antioxidants, which are effective in reducing the risk of heart disease, immune-system decline, different inflammatory processes, etc. [1,2]. Honey species also possess antibacterial activities and are scavengers of active oxygen radicals [3]. Among the components present in honey which are responsible for its antioxidative effect are phenolic compounds (flavonols, flavones, flavonones, benzoic and cinnamic acids). The analysis of phenolic compounds has been regarded as a very promising way of studying the floral and geographical origin of honeys [4,5]. For example, kaempferol has been used as a marker for rosemary honey and quercetin for sunflower honey [6]. Some phenolic acids, such as ellagic acid in heather honey [6] and the hydroxycinnamates (caffeic, p-coumaric and ferulic acids) in chestnut honey have also been used as floral markers [7]. Analysis of phenolic compounds is usually carried out using high-performance liquid chromatography [8–10], although gas chromatography and capillary electrophoresis have also been used in some instances [11,12]. As a detection system, diode-array detec-

夽 Presented at the 25th LC–MS Montreux Symposium, 12–14 November 2008, Montreux, Switzerland. ∗ Corresponding author. E-mail address: [email protected] (M. Biesaga). 0021-9673/$ – see front matter © 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.chroma.2009.07.066

tor has been the most common and HPLC-DAD is the classical and reference method for the analysis of phenolic compounds [13,14]. However, due to the complexity of the honey matrix, it is difficult to obtain sufficient sensitivity for phenolic analysis in real samples. Moreover, the identification of the components that show similar UV–VIS spectra as well as the interpretation of the analytical results is very difficult [10,15]. For this reason, for chromatographic analysis of phenolic compounds in honey, the extensive sample preparation must be performed, particularly for removal of sugars and preconcentration of the analytes. This step is time-consuming and the significant loss of some trace analytes may occur [9,10,16,17]. Mass spectrometry detection has the advantages of providing precise structural information about the eluted compounds since co-elution is not a problem as long as they have different molecular masses. Sensitivity and selectivity of detection can be further increased using tandem MS, when more fragmentation of the precursor and product ions is formed, therefore additional structural information for the identification of phenolic compounds could be obtained. Although HPLC–MS offers a number of key advantages for the analysis of bioactive food components [18], there is no report yet on the use of this coupling for phenolic compounds in honey. Due to the importance of phenolic compounds as well as interest in their identification and quantification, the aim of our study has been to develop the first LC–ESI-MSn method for the determination of three major classes of flavonoids (flavonols, flavones, and flavonones) as well as benzoic and cinnamic acids in honey. The honey samples were diluted by acidified water (pH 2), and analyzed by HPLC–ESI-MS/MS and phenolic compounds were separated on a narrow-bore chromatographic column. Comparison

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between atmospheric pressure chemical ionization (APCI) and electrospray ionization (ESI) was performed by analysis of standards. Fragmentation of analytes for subsequent selective Multiple Reaction Monitoring (MRM) analysis was investigated in negative and positive modes. Sample preparation without removal of sugars and clean-up procedure, followed by fast chromatographic separation allowed the analysis to be completed in a short run time. 2. Experimental 2.1. Reagents The commercial phenolic standards used in this study were: vanillic, gallic, caffeic, syringic, p-coumaric acid, p-hydroxyphenyloacetic acid (p-HPA), chlorogenic acid, ferulic acid, sinapic acid, myricetin, narginin, naringenin, luteolin, apigenin, isorhamnetin as well as quercetin and kaempferol from Sigma (Steinheim, Germany) and p-hydroxybenzoic acid (p-HBA) from POCH (Gliwice, Poland). Rutin was purchased from Merck (Darmstadt, Germany). Methanol was of HPLC gradient grade from Merck (Darmstadt, Germany). Ultrapure water from Milli-Q system (Millipore, Bedford, MA, USA) with a conductivity of 18 MQ was used in all experiments. Stock solutions of phenolic acids were prepared with water, stock solutions of flavonoids with methanol. Diluted mix standards were prepared with water. All solutions were filtered through 0.45 ␮m membranes (Millipore) and degassed prior to use. 2.2. Apparatus The samples of honey were analyzed using Shimadzu liquid chromatographic system consisted of binary pumps LC20-AD, degasser DGU-20A5, column oven CTO-20AC, autosampler SIL20AC, detector UV SPD 20A connected to 3200 QTRAP Mass spectrometer (Applied Biosystem/MDS SCIEX). Compounds were separated on an Atlantis C-18 (50 mm × 2.1 mm, 3 ␮m) column from Waters. As eluent A 2 mM formic acid and eluent B methanol were used. The mobile phase was delivered at 0.2 mL/min in linear gradient mode: 0–3 min 22% B, 10 min 100% B, 12 min 100% B, 13 min 22% B, and 16 min 22% B. A MS system equipped with ESI or APCI operated in negative and positive ion modes. Multiple Reaction Monitoring (MRM) scan mode operated under following conditions: capillary temperature 450 ◦ C, curtain gas pressure at 0.3 MPa, auxiliary gas pressure at 0.3 MPa, discharge current 3 ␮A, ionization mode source voltage: 4.5 kV. Nitrogen was used as curtain and auxiliary gas. For each compound the optimum conditions of MRM were determined in infusion mode (Table 2). Standard solutions were infused into the electrospray source via a 50 ␮m i.d. PEEK capillary using a Harward Apparatus pump at 10 ␮L/min. Continuous mass spectra were obtained by scanning m/z from 50 to 650. 2.3. Sample preparation Honey samples (20 g) were mixed with 100 mL of deionized water adjusted to pH 2 with HCl and stirred in a magnetic stirrer for 15 min. The fluid samples were then filtered through cotton wool to remove the solid particles and then through 0.45 ␮m membrane filters. Although samples contained monosaccharides (glucose + fructose), carmelisation, which could contaminate ion source, was not observed. 3. Results and discussion 3.1. Selection of ionization mode Atmospheric pressure chemical ionization and electrospray ionization have emerged as highly useful methods which allow the direct conjunction with liquid-phase separation techniques [19].

Fig. 1. Mass spectra of naringin (2 mg L−1 ) recorded in: (A) negative mode and (B) positive mode ESI.

ESI is a gentler ionization method than APCI, being less likely to cause the analytes to fragment, and is generally used for a wide range of polar compounds that can be ionized in solution, while APCI is used for less polar molecules that can undergo acid-based reactions in the gas phase. The application of APCI and ESI in both positive and negative mode was evaluated for all compounds. The cone voltage (declustering potential, DP) was optimized in infusion mode with methanol–water (50:50) as the solvent. The obtained spectra are summarized in Table 1 at CE = 50 V. Experimental data show that the fragmentation pathways are independent of ionization mode (APCI or ESI). On the other hand significant differences do occur as regards the relative abundance of various fragment ions. It is in good correlation with previous data [19]. Our study of comparison between APCI and ESI focused on evaluation of measurement sensitivity in analysis of flavonoids. Flavonols and phenolic acids in the positive ionization mode could form many adducts with the cations from the sample or mobile phases, which makes rich fragmentation pattern. The negative mode results in limited fragmentation but provides the highest sensitivity for flavonoids [20,21]. Fig. 1 shows mass spectra of naringin as an example. As it was expected, more ions occur in positive ionization mode and the noise level is relatively high. Flavonoids do not contain nitrogen atoms and have low basicity in the liquid phase. For this reason the formation of protonated molecules [M+H]+ is lower in positive mode than in negative ESI. As could be seen from Fig. 2 higher MRM signal for rutin (glycoside) was observed with ESI in the negative ion mode, which is able to detect this flavonoid at levels as low as 5 ␮g L−1 . In contrast, the limit of detection for APCI (also in the negative mode) was only 10 mg L−1 . For naringenin (aglycone) APCI and ESI gave similar results. Both APCI [22–24] and ESI [20,25–28] appear to be favored

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Table 1 MS/MS characteristics of phenolic compounds. Compound

Ion source

Mr

Ion mode

MS/MS ions m/z (relative abundance, %)

Gallic acid

ESI

170

+ −

127 (100), 153 (89), 109 (83), 107 (67), 81 (75) 125 (100), 79 (8), 81 (6)

+ −

93 (100), 107 (75), 125 (50) 125 (100), 97 (11), 79 (10)

+ −

95 (100), 121 (62), 93 (20) 93 (100)

+ −

111 (100), 121 (40), 95 (58) 93 (100)

+ −

107 (100), 135 (25), 109 (28), 95 (16) 107 (100)

+ −

107 (100) 107 (100), 121 (8)

+ −

163 (100), 89 (38), 135 (22), 117 (21) 191 (100), 85 (18)

+ −

163 (100), 135 (22), 145 (19), 117 (22) 191 (100), 85 (15)

+ −

93 (100), 125 (50) 123 (100), 108 (91), 152 (58)

+ −

93 (100), 125 (5) 108 (100), 108 (83), 123 (75), 152 (98)

+ −

163 (100), 135 (30), 117 (30) 135 (100)

+ −

163 (100), 145 (23), 135 (28), 117 (30), 89 (51) 135 (100), 107 (5), 117 (3)

+ −

140 (100), 155 (60), 125 (43), 95 (22) 182 (100), 121 (90), 153 (59), 167 (50), 138 (30)

+ −

140 (100), 155 (59), 125 (35) 123 (100), 182 (64), 121 (83), 153 (64), 95 (68)

+ −

207 (100), 155 (59), 125 (35) 164 (100), 149 (85), 121 (70)

+ −

207 (100), 175 (38), 119 (25), 91 (30), 147 (23) 93 (100), 121 (90), 208 (87), 164 (85), 149 (80)

+ −

147 (100), 91 (55), 119 (50) 119 (100), 93 (5)

+ −

147 (100), 137 (38), 119 (43), 91 (85), 121 (20) 119 (100), 93 (10)

+ −

177 (100), 145 (56), 117 (50), 89 (46) 134 (100), 178 (22), 149 (15)

+ −

177 (100), 145 (55), 89 (60), 117 (44), 167 (17) 134 (100), 149 (16), 178 (32)

+ −

153 (100), 217 (35), 245 (20) 151 (100), 179 (63), 109 (53), 107 (33)

+ −

153 (100), 217 (36), 246 (36), 273 (30), 165 (34) 151 (85), 179 (95), 137 (100), 109 (53), 107 (33)

+ −

153 (100), 229 (67), 137 (61), 165 (39) 151 (100), 179 (89), 107 (59), 121 (41)

+ −

153 (100), 229 (60), 137 (59), 165 (42) 151 (100), 179 (31), 121 (25), 107 (21)

+ −

153 (100), 119 (50), 91 (48), 121 (19) 117 (100), 151 (30), 121 (7), 107 (22)

+ −

153 (100), 91 (49), 119 (46), 121 (17) 117 (100), 151 (33), 107 (23), 149 (22)

APCI

p-HBA

ESI

138

APCI

p-HPA

ESI

152

APCI

Chlorogenic acid

ESI

354

APCI

Vanillic acid

ESI

168

APCI

Caffeic acid

ESI

180

APCI

Syringic acid

ESI

198

APCI

Sinapic acid

ESI

222

APCI

p-Coumaric acid

ESI

164

APCI

Ferulic acid

ESI

194

APCI

Myricetin

ESI

318

APCI

Quercetin

ESI

302

APCI

Apigenin

ESI APCI

270

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Table 1 (Continued ) Compound

Ion source

Mr

Ion mode

MS/MS ions m/z (relative abundance, %)

Luteolin

ESI

286

+ −

153 (100), 89 (45), 115 (10), 128 (10) 133 (100), 151 (25), 107 (18),

+ −

153 (100), 135 (50), 117 (25), 161 (20) 133 (100), 151 (24), 107 (15), 175 (13)

+ −

153 (100), 121 (65), 165 (40), 213 (23), 128 (20) 151 (100), 93 (45), 117 (30), 145 (25)

+ −

153 (100), 121 (59), 165 (32), 93 (26) 151 (100), 93 (45), 107 (38), 108 (35)

+ −

153 (100), 147 (48), 91 (26), 119 (23) 151 (100), 119 (97), 107 (39), 83 (18), 177 (15)

+ −

153 (100), 147 (54), 91 (27), 119 (27) 119 (100), 151 (68), 107 (33), 83 (18)

+ −

273 (100), 153 (55), 85 (23), 147 (15) 151 (100), 271 (65), 119 (49), 107 (34), 459 (8)

+ −

273 (100), 153 (55), 147 (15), 119 (10) 151 (100), 271 (58), 119 (41)

+ −

303 (100), 465 (15), 85 (15), 153 (5) 300 (100), 271 (48), 301 (37), 255 (23), 151 (15)

+ −

303 (100), 85 (21), 153 (5) 300 (100), 271 (45), 301 (28), 255 (24)

APCI

Kaempferol

ESI

286

APCI

Naringenin

ESI

272

APCI

Naringin

ESI

580

APCI

Rutin

ESI APCI

610

Fig. 2. MRM signals of (A) rutin (10 mg L−1 ) and (B) naringenin (2 mg L−1 ) with APCI and ESI negative ionization modes at optimal conditions after chromatographic separation.

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Table 2 LC/MS/MS characteristics of phenolic compounds in the negative mode. Compound

Retention time (min)

MRM

DP (V)

CE (V)

Gallic acid p-HPA p-HBA Chlorogenic acid Vanillic acid Caffeic acid Syringic acid p-Coumaric acid Ferulic acid Sinapic acid Rutin (quercetin-3-O-rutnoside) Myricetin Naringin (naringenine-7-O-rhamonoglucoside) Naringenin Quercetin Luteolin Kaempferol Apigenin Isorhamnetin

0.85 1.6 1.7 2.0 2.1 2.3 2.6 3.9 4.5 5.5 6.7 7.0 7.5 7.6 7.7 7.9 8.1 8.3 8.9

169/125 151/107 137/93 353/191 167/152 179/135 197/182 163/119 193/134 223/121 609/301 317/179 579/271 271/151 301/151 285/133 285/151 269/117 315/199

−45 −15 −25 −20 −30 −10 −20 −20 −30 −35 −80 −55 −70 −35 −70 −80 −70 −50 −55

−20 −12 −18 −22 −16 −24 −24 −18 −20 −38 −48 −34 −56 −26 −30 −44 −36 −46 −32

by different authors for the analysis of flavonoids. However, the eluent composition could have a significant influence on the ionization efficiency. The eluent combination methanol–ammonium formate at pH 4.0 gave the highest response in APCI (negative

mode) [23], while ESI-MS with an acidic ammonium acetate buffer and the organic component consisted of methanol and acetonitrile as the mobile phase provides the best sensitivity [25]. Moreover, analyte response could vary considerably from one sub-class of

Fig. 3. Source fragmentation of (A) ferulic acid (1 mg L−1 ) and (B) naringenin (1 mg L−1 ) in negative ESI at different collision energy.

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phenolic compounds to another, and even within one class [19,25]. The results reported by Cluyckens and Claeys [28] agree with our findings. Although the linear range in APCI mode is usually wider compared to ESI, quantitation in ESI might be advantageous when working with small concentrations. 3.2. Mass spectra The spectra in negative ESI were dominated by [M−H]− ions and these ions were selected for collision-activated dissociation studies of the precursor ions. It has been demonstrated that optimization of the ESI parameters plays a key role in the achievement of adequate MS signals for any analyte [29,5]. The declustering potential (DP) and collision energy (CE), measured as fragmentor voltage value, were optimized in infusion mode for each compound in the range from −400 to −10 V and −130 to −5 V, respectively. The voltage required for significant fragmentation was compound dependent parameter (Table 2). The product ion spectrum of ferulic acid at different collision energies is shown in Fig. 3A. Except the deprotonated molecule [M−H]− , some fragments could be seen even at relatively low CE. Loss of CO2 was observed giving the [M−H44]− (m/z 149) as a characteristic ion and the loss of the methyl group, providing a [M−H-15]−• anion radical at m/z 178. As collision energy was increased the intensity of both peaks was reduced in favor of the [M−H-44-15]− (m/z 134) peak. For most phenolic acids the MRM method was built on the loss of a one methyl group [M−H−CH3 ]− (vanilic, syringic, ferulic), or carboxylic group [M−H−CO2 ]− (p-HBA, gallic, caffeic, p-coumaric). The studied phenolic acids contain two distinct acidic groups, carboxylate and phenolic hydroxyls. The deprotonation site is probably the more acidic of these two but deprotonation of the phenolic hydroxyl is also possible [30]. However, there were no doubly charged ions in the spectra. Chlorogenic acid showed characteristic product ion m/z 191 which corresponds to the deprotonated quinic acid. Fragmentation patterns of flavonoids were also studied at different collision energy. Fig. 3 shows the obtained results for ferulic acid and naringenin in negative ESI. In the spectra of flavonoids, the only ions detected at low collision energy were the deprotonated molecules (Fig. 3B). The glycosidic bond is weaker than the bonds within the flavonoid molecule, thus the fragmentation pathway of O-glycolylated flavonoids typically starts with the cleavage of this bond, leading to elimination of the sugar moiety with the charge being retained on the aglycone fragment. Higher collision activation led to sequential losses of sugar moiety of 308 Da from naringin and rutin. For aglycons the base peaks arise from Retro-Diels–Alder reactions providing information on the number and type of substituents in the A- and B-rings. These fragmentation pathways allow the distinction between luteolin (flavone) and kaempferol (flavonol) although both flavonoids have the same molecular mass. The conditions for transitions for all tested compounds were collected in Table 2. 3.3. Optimization of LC–MS method The LC-DAD method previously used [10] was modified to be compatible with the LC–MS system. The shorter column and small particles (50 mm × 2.1 mm, 3 ␮m) allowed the separation of all 19 compounds studied in less than 9 min using gradient elution. The elution sequence is directed by their structural characteristics. Generally, the retention times followed the expected reversed-phase pattern of benzoic acids < cinnamic acids < flavonoid O-glycosides < flavonoid aglycones (Table 2). Among flavonoids, hydroxylation decreases retention owing to increasing polarity (hydrogen bond formation ability) and the elution pattern is affected by the number of OH-groups—myricetin has three OH-groups at 3 , 4 and 5 position in the B-ring and it

Fig. 4. Total ion current of MRM of buckwheat honey extract and traces of phenolic compounds in MRM mode. MS/MS conditions described in the text.

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Table 3 Content (mg/kg) of phenolic compounds in honey samples (n = 3). Compound

Buckwheat

Acacia

Gallic acid p-HPA p-HBA Chlorogenic acid Vanillic acid Caffeic acid Syringic acid p-Coumaric acid Ferulic acid Rutin Myricetin Naringin Naringenin Quercetin Apigenin

– 0.7 ± 22.6 ± 0.5 ± 0.4 ± 1.0 ± 0.5 ± 11.5 ± 54.4 ± 0.6 ± 0.2 ± 0.4 ± 0.09 ± 0.4 ± 1.6 ±

0.2 1.7 3.9 0.5 0.5 0.7 0.7 2.2 139.8 0.5 0.1 0.3 0.3 1.2 0.6

0.05 0.68 0.04 0.03 0.04 0.03 0.4 2.2 0.04 0.01 0.02 0.004 0.03 0.03

Honeydew ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.03 0.07 0.12 0.04 0.03 0.02 0.06 0.09 5.6 0.03 0.01 0.01 0.03 0.06 0.05

0.3 ± 4.3 ± 1.6 ± 0.5 ± 0.4 ± 0.7 ± 0.7 ± 2.7 ± 173.8 ± 0.4 ± 0.1 ± 0.3 ± – 0.1 ± 0.3 ±

0.03 0.13 0.03 0.03 0.02 0.05 0.04 0.09 8.5 0.02 0.02 0.02 0.01 0.02

is eluted as first in this group of compounds. Methylation of the hydroxy group in ring B (as in the case of isorhamnetin) causes slightly increased retention. The linearity of the detector response was determined by the square correlation coefficients of the calibration curves generated by three repeated injections of standard solutions at six concentration levels (0.05–50 mg L−1 ). All the compounds showed a good linearity with regression coefficients ≥ 0.996. Limits of detection (LODs) were estimated by decreasing the concentration of the analyte down to the smallest detectable peaks and then this concentration was multiplied by three. LODs were ranged between 1 and 15 ng L−1 for p-coumaric acid and naringenin, respectively. Since the values for the limit of detection and quantification are low in the relation to the usual content of phenolic compounds in honey samples [31], it could reasonable to conclude that this method could be used for reliable quantitative analysis without preconcentration step. The precision was established by assaying six different extracts of the same honey sample with the proposed chromatographic analysis. The RSD were in the range of 1.3–6.8% for the studied compounds. 3.4. Characterization of honey phenolic compounds The chromatogram of the extract of buckwheat honey by LC/MS/MS is presented in Fig. 4. Peak identity was established by both the retention time compared to that of authentic standards and the characteristic transitions (precursor and product ion pair). The traces for p-HBA, p-HPA, caffeic acid, p-coumaric acid, quercetin and apigenin, respectively, found in honey extract in the MRM mode (Fig. 4). This approach offers the possibility of rapid profiling using MS and MS/MS to monitor product ion or selected fragments. In our previous paper [10] determination of p-coumaric acid was impossible using LC-DAD even after clean-up step with an SPE column packed with Oasis HLB sorbent. In the extracts of linden and heather honey samples, peaks at the same retention time of this analyte were observed, but its absorbance spectra were different from that corresponding to standard solution. The results of the quantitative determination of phenolic acids and flavonoids in honey samples from different botanical origin are presented in Table 3. Comparison with literature values is difficult since previous studies [8,9,11] have employed different conditions

for sample preparation to those used here. Moreover, different polyphenolic acids as well as the kind of flavonoid compounds were determined [12,17,18]. The data concerning the content of phenolic compounds in buckwheat honey are not available. This kind of honey tends to exhibit higher content of p-HBA, p-coumaric acid and apigenin relatively to those found in acacia and honeydew. To the contrast, ferulic acid presented lower content. The levels of pHPA, caffeic and syringic acids were in agreement with previous studies on acacia [8] and honeydew samples [32]. Acknowledgments The authors would like to thank the Structural Research Laboratory (SRL) at the Department of Chemistry of University of Warsaw for using HPLC–MS. SRL has been established with financial support from European Regional Development Found in the Sectorial Operational Programme “Improvement of the competitiveness of Enterprises, years 2004–2005” project no: WPK 1/1.4.3./1/2004/72/72/165/2005/U. This work was also partially financed by 501/68-BW-172101 project. References [1] A.M. Gómez-Carcaca, M. Gómez-Romero, D. Arráez-Román, A. SeguraCarretero, A. Fernández-Gutiérrez, J. Pharm. Biomed. Anal. 41 (2007) 1220. [2] M. Blasa, M. Candiracci, A. Accorsi, M.P. Piacentini, E. Piatti, Food Chem. 104 (2007) 1635. [3] T. Nagai, R. Inoue, N. Kanamori, N. Suzuki, T. Nagashima, Food Chem. 97 (2006) 256. [4] M. Küc¸ük, S. Kolayh, S. Karao˘glu, E. Ulusoy, C. Baltaci, F. Candan, Food Chem. 100 (2007) 526. [5] L.F. Cuevas-Glory, J.A. Pino, L.S. Santiago, E. Sauri-Duch, Food Chem. 103 (2007) 1032. [6] F.A. Thomás-Barberán, I. Martos, F. Ferreres, B.S. Radovic, E. Anklam, J. Sci. Food Agric. 81 (2001) 485. [7] S.M. Antony, I.Y. Han, J.R. Rieck, P.L. Dawson, J. Agric. Food Chem. 48 (2000) 3985. [8] L. Yao, Y. Jiang, B. D‘Arcy, R. Singanusong, N. Datta, N. Caffin, K. Raymont, J. Agric. Food Chem. 52 (2004) 210. [9] B. Dimitrova, R. Gevrenova, E. Anklam, Phytochem. Anal. 18 (2007) 24. [10] A. Michałkiewicz, M. Biesaga, K. Pyrzynska, J. Chromatogr. A 1187 (2008) 18. [11] E. De la Fuente, I. Martinez-Castro, J. Sanz, J. Sep. Sci. 28 (2005) 1093. [12] D. Arráez-Román, A.M. Gómez-Carcaca, M. Gómez-Romero, A. SeguraCarretero, A. Fernández-Gutiérrez, J. Pharm. Biomed. Anal. 41 (2007) 1648. [13] H.N. Merken, G.R. Beecher, J. Agric. Food Chem. 48 (2000) 577. [14] R. Tsao, Z. Deng, J. Chromatogr. A 812 (2004) 85. [15] A.M. Gómez-Carcaca, A. Segura-Carretero, A. Fernández-Gutiérrez, Agro. Food Ind. Hi-tech. 17 (2006) 68. [16] L. Martos, F. Ferreres, F.A. Tomás-Barberán, J. Agric. Food Chem. 48 (2000) 1498. [17] S. Suárez-Luque, I. Mato, J.F.J.F. Huidobro, J. Simal-Lozano, J. Chromatogr. B 770 (2002) 77. [18] K. Robards, J. Chromatogr. A 1000 (2003) 657. [19] E. dr Rijke, P. Out, W.M.A. Niessen, F. Ariese, C. Gooijer, U.A.Th. Brinkman, J. Chromatogr. A 1112 (2006) 31. [20] M. Pikulski, A. Aquilar, J.S. Brodbelt, J. Am. Soc. Mass Spectrom. 18 (2007) 422. [21] L. Wang, M.E. Morris, J. Chromatogr. B 821 (2005) 194. [22] U. Justesen, J. Chromatogr. A 902 (2000) 369. [23] E. de Rijke, H. Zappey, F. Ariese, C. Gooijer, U.A.Th. Brinkman, J. Chromatogr. A 984 (2003) 45. [24] S.M. Boué, C.H. Carter-Wientjes, B.Y. Shih, T.E. Cleveland, J. Chromatogr. A 991 (2003) 61. [25] J.P. Rauha, H. Vuorela, R. Kostiainen, J. Mass Spectrom. 36 (2001) 1269. [26] W. Wu, C. Yan, L. Li, S. Li, J. Chromatogr. A 1047 (2004) 213. [27] R.E. March, E.G. Lewars, C.J. Stadey, X.S. Miao, X. Zhao, C.D. Metcalfe, Int. J. Mass Spectrom. 248 (2006) 61. [28] F. Cluyckens, M. Claeys, Rapid Commun. Mass Spectrom. 24 (2002) 2341. [29] W. Mullen, A. Boitier, A.J. Steward, A. Crozier, J. Chromatogr. A 1058 (2004) 163. [30] P. Swatsitang, G. Tucker, K. Robards, D. Jardine, Anal. Chim. Acta 417 (2000) 233. [31] P. Mattila, J. Hellström, J. Food Comp. Anal. 20 (2007) 252. [32] R.J. Weston, L.K. Brocklebank, Y. Lu, Food Chem. 70 (2000).