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Identification and quantitative analysis of β-sitosterol oxides in vegetable oils by capillary gas chromatography–mass spectrometry
Steroids 70 (2005) 896–906
Identification and quantitative analysis of -sitosterol oxides in vegetable oils by capillary gas chromatography–mass spectrometry Xin Zhang a , Diane Julien-David a , Michel Miesch b , Philippe Geoffroy b , Francis Raul c , Stamatiki Roussi c , Dalal Aoud´e-Werner d , Eric Marchioni a,∗ a
b
Laboratoire de Chimie Analytique et Sciences de l’Aliment (UMR 7512), Facult´e de Pharmacie, Universit´e Louis Pasteur, 74 route du Rhin, 67400 Illkirch, France Laboratoire de Chimie Organique Synth´etique (UMR 7123), Universit´e Louis Pasteur, 1 rue Blaise Pascal, 67008 Strasbourg, France c Laboratoire d’Oncologie Nutritionnelle, Inserm UMR S392/IRCAD, 1 place de l’Hˆ opital, 67091 Strasbourg, France d A´ erial, rue Laurent Fries—Parc d’Innovation, 67412 Illkirch, France Received 28 October 2004; received in revised form 9 June 2005; accepted 9 June 2005 Available online 20 July 2005
Abstract As vegetable oils and phytosterol-enriched spreads are marketed for frying food or cooking purposes, temperature is one of the most important factors leading to the formation of phytosterol oxides in food matrix. A methodology based on saponification, organic solvent extraction, solid-phase extraction (SPE), followed by mass spectrometric identification and quantitation of -sitosterol oxides using capillary gas chromatography–mass spectrometry (GC–MS) in selected ion monitoring (SIM) mode was developed and characterized. Relative response factors of six -sitosterol oxides, including 7␣-hydroxy, 7-hydroxy, 5,6␣-epoxy, 5,6-epoxy, 7-keto, and 5␣,6-dihydroxysitosterol, were calculated against authentic standards of 19-hydroxycholesterol or cholestanol. Linear calibration data, limit of detection, and sample recoveries during analytical process. Recoveries of these oxidation compounds in spiked samples ranged from 88 to 115%, while relative standard derivation (R.S.D.) values were below 10% in most cases. The analytical method was applied to quantify -sitosterol oxides formed in thermal-oxidized vegetable oils which were heated at different temperatures and for varying time periods. Sitosterol oxidation is strikingly higher in sunflower oil relative to olive oil under all conditions of temperature and heating time. © 2005 Elsevier Inc. All rights reserved. Keywords: -Sitosterol oxides; Gas chromatography–mass spectrometry (GC–MS); Selected ion monitoring (SIM) mode; Quantification; Oil; Thermaloxidation
1. Introduction Phytosterols in edible vegetable oils or plant sterolenriched foods are structurally similar to cholesterol, the only differences being the position of double bond and the alkyl side chain. Both cholesterol and phytosterols contain an unsaturated ring-structure and are thus susceptible undergo oxidation in the presence of oxygen. Oxidation is accelerated by heating, exposure to ionizing radiation, light, chemical catalysts, or enzymatic processes [1–5]. For cholesterol, more than 80 oxides have been identified and extensively ∗
Corresponding author. Tel.: +33 3 90244326; fax: +33 3 90244325. E-mail address: [email protected] (E. Marchioni).
0039-128X/$ – see front matter © 2005 Elsevier Inc. All rights reserved. doi:10.1016/j.steroids.2005.06.004
studied and reviewed [6–9]. The main oxides are hydroxy, keto, and epoxy compounds. The latter may hydrate to dihydroxy sterols [6]. The in vitro studies have shown that some cholesterol oxides have cytotoxic, mutagenic, atherogenic, and carcinogenic activities [10,11]. Due to the structural similarity with cholesterol and its oxides, phytosterols and their oxides have received much attention during the last decades with respect to their biological properties and consequently, their performance in food safety evaluation [12–16]. Some studies [9,12] have shown that supplementation of foods with phytosterols and phytosterol esters may induce the presence of their oxides in foods and human plasma. The analytical methods for phytosterol oxides are based on those developed for cholesterol oxides. Gas chromatography
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(GC) or high performance liquid chromatography (HPLC) is most often employed for the separation and quantification of cholesterol oxides. Although HPLC provides good resolution for cholesterol oxides [1,17], this technique is not suitable for phytosterol oxides analysis because of excessive interference, modest sensitivity, and limited chromatographic resolution. For this reason, final separation and identification of phytosterol oxides in food matrix are always carried out by GC–FID or GC–MS [18–24]. Guardiola et al. [25] and Diczfalusy [26] reviewed the analysis of sterol oxides in foods and biological samples. Studies dealing with sterol oxide mixtures from complex food samples using GC–MS operating in selected ion monitoring (SIM) mode as quantitative technique may increase the detection selectivity and avoid the numerous interfering problems [27,28]. Since individual standards of phytosterol oxides are not available commercially, most research works, including chemical quantitative analysis, biochemical, and medical studies, have used mixtures of phytosterol oxides [15,16,20]. Six individual -sitosterol oxides have been synthesized recently [29]. In the present study, these compounds were used as standards, a precise quantitative analytical method of these products formed in thermal-oxidized oils are performed. Much attention was paid to the formation of oxides when heating vegetable oils for short periods of time (≤1 h).
2. Experimental 2.1. Materials and reagents 5␣-Cholestan-3-ol (cholestanol, 95%) and cholest-5ene-3,19-diol (19-hydroxycholesterol, 95%), purchased from Sigma (Steinheim, Germany), were used as internal standards. -Sitosterol was purified from Generol 95R (Cognis, Saint-Fargeau-Ponthierry, France), a commercial phytosterol mixture, in the laboratory [29]. -Sitosterol oxides, e.g., sitost-5-ene-3,7␣-diol (7␣-hydroxysitosterol), sitost-5-ene-3,7-diol (7-hydroxysitosterol), 5,6␣-epoxy5␣-sitostan-3-ol (5,6␣-epoxysitosterol), 5,6-epoxy-5sitostan-3-ol (5,6-epoxysitosterol), 3-hydroxysitost-5en-7-one (7-ketositosterol), and sitost-5-en-3,5␣,6-triol (5␣,6-dihydroxysitosterol), were then chemically synthesized [29]. The purity of each compound was ∼95% as determined by 1 H NMR, 13 C NMR, and GC–MS. Stock solution (1 mg/mL) of each analyte and internal standards, cholestanol and 19-hydroxycholesterol, were prepared separately in ethyl acetate and stored at 4 ◦ C in the dark. Ethyl acetate was obtained from Acros (Geel, Belgium), ethanol from Carlo Erba (France), cyclohexane from Fluka (Buchs, Switzerland), diethyl ether from Riedel-de Ha¨en (Seelze, Germany), acetone and pyridine from Prolabo (Fontenay sous Bois, France), isooctane and potassium hydroxide (KOH) from Merck (Darmstadt, Germany), and anhydrous sodium sulfate (Na2 SO4 ) from SDS (Peypin, France). All organic solvents and reagents were
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of analytical grade. Ultra-pure water was obtained by means of a Millipore Q water purification system (Milford, MA, USA). N-Methyl-N-(trimethylsilyl)trifluoroacetamide (MSTFA), used as the silylation reagent, was provided by Sigma-Aldrich (Steinheim, Germany). The SiOH-SPE cartridges (3 mL/500 mg) were purchased from MachereyNagel (Chromabond® , D¨uren, Germany). 2.2. Food samples Sunflower oil (1 L, Lesieur Tournesol, France), olive oil (0.5 L, Puget, France), and butter (250 g, Pr´esident, France) were of edible quality and purchased in a local supermarket. 2.3. Samples pretreatment Firstly, 1 mg of -sitosterol was weighed in an open glassware (10 mm × 60 mm i.d.) and heated in an oven (Memmert, Schwabach, Germany) which had been set at 100, 150, or 200 ◦ C for 5, 15, and 30 min. The final products were converted to TMS ethers (as described below) and analyzed directly by GC–MS to estimate the main oxides formed. After the model assay, approximately 200 mg of oil samples (sunflower oil or olive oil) was weighted in the same type of glassware to ensure efficient surface contact with air, and then, they were placed into the oven and heated at 150 or 200 ◦ C. After various durations (from 5 to 60 min), the glasswares were removed from the oven, quickly covered and sealed with aluminium foil, and allowed to cool to room temperature. The samples were then dissolved in excess ethyl acetate, collected into 30 mL brown vials, evaporated to dryness under a gentle nitrogen flow, and stored at 4 ◦ C under nitrogen. Each sample was assayed in triplicate. 2.4. Isolation and purification of phytosterol oxides from food samples The procedure for sample preparation described here for phytosterol oxides analysis was adapted from the methods proposed by Dutta [18] and Lampi et al. [20] with minor changes. The detailed processes were as follows: • Cold saponification. Prior to saponification, 20 L of 19hydroxycholesterol solution (1 mg/mL in ethyl acetate) as an internal standard was spiked into the heated 200 mg of oil samples in a 30 mL brown vial. After removal of the ethyl acetate under nitrogen, 9 mL of ethanol, and 0.5 mL of saturated KOH aqueous solution were added into the vial. The sample was purged with nitrogen, and the tightly closed brown vial was put into a rotary shaker (Edmund B¨uhler, Johanna Otta GmbH, Hechingen, Germany) at ambient temperature in the dark overnight (15 h). • Extraction. The solution was transferred in a 100 mL separation funnel and 10 mL of distilled water were added. The unsaponifiable fraction was extracted with diethyl ether (4× 15 mL). The combined organic extracts were
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washed with 20 mL of 0.5 M aqueous KOH solution and 0.2 M aqueous Na2 SO4 solution (3× 15 mL). The organic solvents were removed with a rotary vacuum evaporator (25 ◦ C, 200 mbar), and the residue was dried under nitrogen flow. After additional 2 mL of diethyl ether, the unsaponifiable residue was carefully transferred using a pipette to a 2 mL glass test tube and then evaporated to dry matter under nitrogen flow. • Solid-phase extraction (SPE) purification. The SiOH-SPE cartridges were first conditioned with 5 mL of cyclohexane. The unsaponifiable matter was dissolved in 1 mL of a solution of cyclohexane/diethyl ether (9:1, v/v) and was loaded onto the cartridge. Low-polarity lipids and non-oxidized sterols were eluted with 5 mL of cyclohexane/diethyl ether (9:1, v/v) followed by 4 mL of cyclohexane/diethyl ether (1:1, v/v). Finally, the sterol oxides were eluted out of the cartridge with 5 mL of acetone and collected into a test tube to which 20 L of cholestanol (1 mg/mL in ethyl acetate) were added. This residue was dried under a gentle nitrogen flow. • Derivatization. The purified sterol oxides residue was converted to trimethylsilyl (TMS) ethers with 50 L of pyridine and 40 L of MSTFA at room temperature in the dark overnight and then, diluted with 410 L of isooctane. One microliter of each sample was injected into the GC–MS system. 2.5. Gas chromatography–mass spectrometry (GC–MS) analysis GC–MS analyses were performed on a Varian STAR 3400 GC instrument equipped with an on-column SPI injector coupled to a Varian SATURN 2000 mass sensitive detector (Varian, France). Data acquisition and processing, and instrumental control were performed by Varian Saturn WS software. Analytes were separated in a VF-5ms capillary column (phase stationary: 5% phenyl–95% dimethylpolysiloxane, thickness of 0.1 m, 60 m × 0.25 mm, Varian, France). The column temperature gradient was programmed from 105 ◦ C (hold for 2 min) to 170 ◦ C at 20 ◦ C/min and then, to 320 ◦ C at 7 ◦ C/min (hold for 15 min). The injector operating conditions were as follows: injection volume 1 L; initial injector temperature of 105 ◦ C was increased to 300 ◦ C at 100 ◦ C/min (hold for 40 min). Helium (purity 99.9995%) was used as a carrier gas with a flow rate of 1 mL/min. Electronic impact (EI) ionization mode mass spectra were obtained at 70 eV and monitored on the full-scan range (m/z 40–600). Quantitative data were carried out using selected ion monitoring (SIM) analysis. The analytes were quantified as follows: Ac τis nc = nis RRFc Ais τc where nc and nis were the moles of component analyzed in the sample and of internal standard added to the sample, respectively; Ac and Ais the peak areas of component analyzed in
the sample and of internal standard added to the sample, respectively; τ c and τ is the method recoveries of component analyzed in the sample and of internal standard added to the sample, respectively; and RRFc was the relative response factor of each pure -sitosterol oxide compared to the internal standard, which was calculated as follows: RRFc =
nc Ais RRFis nis Ac
where Ac and Ais were the peak areas of standard oxide and of internal standard, respectively; nc and nis the moles of standard oxide and of internal standard, respectively; and RRFis was the response factor of the internal standard, which was set to 1. 2.6. Linearity, limit of detection and recovery For each oxide, standard solutions were made at six different concentrations between 0.1 and 100 g/mL, derivatized, and assayed in duplicate. Linear plots of peak areas versus concentration were calculated. Recoveries of SiOH-SPE purification were tested with mixtures containing the six sitosterol oxides as well as 19-hydroxycholesterol (10, 20, and 40 g of each one) spiked with 20 g of cholestanol as an internal standard. Recoveries of the complete analytical method were carried out in 200 mg of butter spiked with the six -sitosterol oxides (10, 20, and 40 g) and 20 g of 19hydroxycholesterol as an internal standard.
3. Results and discussion 3.1. Identification and characterization of β-sitosterol oxides by GC–MS A series of individual -sitosterol oxides (∼95% purity), including 7␣-hydroxy, 7-hydroxy, 5,6␣-epoxy, 5,6-epoxy, 7-keto, and 5␣,6-dihydroxysitosterol, were synthesized from purified -sitosterol and characterized by means of GC–MS as TMS ethers. Chromatographic separations of these compounds were well achieved on a VF-5ms capillary column and identified by their MS data. Retention times were consistent within 1.2 s across runs. The typical retention times and selected ions chosen for identification and quantitation along with their relative abundances are summarized in Table 1. The mass spectra of these oxides showed the similar typical fragmentation patterns, which were in agreement with the data reported previously [2,30,31] with minor differences in the relative abundances. One point should be mentioned that some mass spectra rounded numbers to the unit, giving occurrence to impaired mass fragments of molecular ion or base peak. For the mass spectra of 7␣-hydroxy and 7-hydroxysitosterol as di-TMS ether derivatives, the base peak was observed at m/z 485 (M+ − TMSOH). The relative intensity of molecular ions (M+ ) at m/z 575 was 1%.
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Table 1 Retention time (tR ) and relative retention time (RRT) of -sitosterol and its oxides as TMS ethers in relation to that of 19-hydroxycholesterol, as well as their specific ions Analytes
tR (min)
RRTa
Mw b
SIM ions (relative abundance, %) Identification
Quantitation
-Sitosterol Cholestanol 19-Hydroxycholesterol 7␣-Hydroxysitosterol 7-Hydroxysitosterol 5,6-Epoxysitosterol 5,6␣-Epoxysitosterol 5␣,6-Dihydroxysitosterol 7-Ketositosterol
31.42 29.28 29.55 30.48 32.03 33.08 33.32 34.07 35.58
1.06 0.99 1.00 1.03 1.08 1.11 1.12 1.15 1.20
486.7 460.7 546.7 574.7 574.7 502.7 502.7 664.7 500.7
382 (52), 397 (100), 472 (16), 487 (32) 215 (100), 356 (65), 446 (29), 461 (12) 73 (100), 353 (75), 367 (59), 547 (3) 470 (4), 485 (100), 575 (1) 470 (3), 485 (100), 575 (1) 73 (100), 395 (38), 413 (52), 503 (24) 73 (100), 395 (60), 413 (46), 503 (15) 432 (100), 485 (84), 560 (34), 575 (13) 396 (100), 486 (41), 501 (80)
397 356 353 485 485 413 395 432 396
a b
Relative to the retention time of 19-hydroxycholesterol TMS ether. Mw : molecular weight as TMS ether.
The other typical fragments 470 (M+ − TMSOH-CH3 ), 395 (M+ − 2TMSOH), 253 (M+ − 2TMSOH-side chain), and 129 (5-sterol) were of low intensity (1–6%), except for m/z 73 (22%). For the TMS ether derivatives of 5,6␣-epoxy and 5,6-epoxysitosterol, similar ion fragmentation patterns were found, apart from some differences in their relative peak intensities. The molecular ion at m/z 503 was present for both compounds (15 and 24%), and the base peak was at m/z 73. The molecular ion of 7-ketositosterol TMS ether was at m/z 501 (M+ , 80%), and the base peak was at m/z 396 (M+ − TMSOH-CH3 , 100%). This result was different from previously reported data [28,31] in which an unspecified base peak at m/z 174 was observed. The other typical fragments were consistent with the data reported previously [31,32]. Special attention should be taken with 5␣,6dihydroxysitosterol. Indeed, depending on the silylating reagents and on the reaction time used, bis- and tris-TMS ether derivatives may be produced from 5␣,6dihydroxycholesterol or 5␣,6-dihydroxysitosterol because the derivatization at the 5␣-hydroxyl group of the sterol is sterically hindered [31,32]. In the present work, when derivatization was performed at 50 ◦ C for 60 min, three
hydroxyl groups of 5␣,6-dihydroxysitosterol could not completely be derivatized to the tris-TMS ether derivative. A gas chromatogram showed two peaks, one (tR = 33.98 min) corresponding to tris-TMS ether (with three OTMS groups) and the other (tR = 37.21 min) to the bis-TMS ether (with one hydroxy and two OTMS groups) (Fig. 1(a)). To avoid this phenomenon, the derivation reaction between analytes and MSTFA was carried out at ambient temperature overnight in the dark. The three hydroxyl groups of 5␣,6-dihydroxysitosterol could then be converted to tris-TMS ether (Fig. 1(b)). Its base peak was observed at m/z 432 (M+ − TMSOH-side chain). The other major fragments as well as their relative intensities were at m/z 575 (M+ − TMSOH, 13%), 560 (M+ − TMSOH-CH3 , 34%), 485 (M+ − 2TMSOH, 84%), 470 (M+ − 2TMSOH-CH3 , 28%), 395 (M+ − 3TMSOH, 11%), and 380 (M+ − 3TMSOH-CH3 , 4%). Moreover, this method was suitable for the silylation of all other studied oxides present in the samples prior to GC analysis. Relative response factors (RRF) of six -sitosterol oxides were calculated with SIM mode in the present work in relation to the internal standard, cholestanol or 19hydroxycholesterol, are listed in Table 2.
Table 2 Relative response factors (RRF) of -sitosterol and its oxides as TMS ethers in relation to those of cholestanol or 19-hydroxycholesterola Analytes
-Sitosterol Cholestanol 19-Hydroxycholesterol 7␣-Hydroxysitosterol 7-Hydroxysitosterol 5,6-Epoxysitosterol 5,6␣-Epoxysitosterol 5␣,6-Dihydroxysitosterol 7-Ketositosterol a b c
RRF (SIM mode)c
RRF (TIC mode) Relative to cholestanol
Relative to 19hydroxycholesterol
Relative to cholestanol
Relative to 19hydroxycholesterol
1.06 ± 0.04b – 0.69 ± 0.05 0.79 ± 0.05 0.85 ± 0.07 1.99 ± 0.31 1.25 ± 0.09 0.62 ± 0.29 1.31 ± 0.09
– 1.45 ± 0.08 – 1.05 ± 0.09 1.18 ± 0.08 2.73 ± 0.17 1.66 ± 0.14 0.86 ± 0.07 1.75 ± 0.13
0.17 ± 0.01b – 0.77 ± 0.03 0.12 ± 0.01 0.13 ± 0.01 6.44 ± 0.20 3.54 ± 0.08 0.35 ± 0.02 1.00 ± 0.08
– 1.29 ± 0.05 – 0.16 ± 0.01 0.17 ± 0.01 8.29 ± 0.28 4.63 ± 0.16 0.49 ± 0.02 1.31 ± 0.12
Mean ± standard derivation (n = 10). Mean ± standard derivation (n = 3). The characteristic ions for quantitation in SIM mode, see Table 1.
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Fig. 1. Influence of silylation conditions on the derivatization of 5␣,6-dihydroxysitosterol: (a) 50 ◦ C for 60 min, (b) room temperature for overnight. The first silylation condition lead to the formation of two peaks as shown in GC chromatogram (a). The upper left mass spectrum corresponds to tris-TMS ether (peak 1, with three OTMS groups). The upper right mass spectrum corresponds to bis-TMS ether (peak 2, with one hydroxy and two OTMS groups).
In previous works, because the phytosterol oxides were not commercially available, the RRF values of these compounds were usually adopted from those of the corresponding cholesterol oxides [19,20] or assumed to be a value of 1 [33,34] in GC–FID or GC–MS analysis with total ion chromatogram (TIC) mode. However, although good separation and analytical methods had been reported for cholesterol oxides [6,7,9], the chromatographic resolution problems associated with the analysis of phytosterol oxides in foods of plant origin or mixed origin (animal/plant-based foods) are quite complex because of the high probabilities of co-elution [9,22]. Such as, in the present GC separation system, 5,6-epoxysitosterol was co-eluted with 7-hydroxycampesterol. Therefore, due to high relative intensity of some characteristic ions, SIM mode has advantages in sensitivity, identification and quantitation of sterol oxides in food matrix. Guardiola et al. [25]
reviewed the characteristic ions (m/z) used to quantify cholesterol oxides as trimethylsilyl ether derivatives by GC–MS. However, in relation to phytosterol oxides, the literature is scarce. The only work that has quantified 7-ketositosterol in food samples by GC–MS operated in SIM mode was performed by Turchetto et al. [28], who selected the characteristic ions at m/z 500, 484, 410, and 395 to quantify the TMSE derivative of this compound. Since six -sitosterol oxides were synthesized and characterized recently, their exact RRF values were available. The ions selected for quantification were m/z 353 for 19-hydroxycholesterol, m/z 485 for 7␣hydroxy and 7-hydroxy, m/z 413 for 5,6-epoxy, m/z 395 for 5,6␣-epoxy, m/z 432 for 5␣,6-dihydroxy, and m/z 396 for 7-ketositosterol. In some cases, their base peaks were chosen (7␣-hydroxy, 7-hydroxy, 5␣,6-
X. Zhang et al. / Steroids 70 (2005) 896–906
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Table 3 Analytical parameters of gas chromatography for each -sitosterol oxide as TMS ethers assayed by the present method Analytes
19-Hydroxycholesterol 7␣-Hydroxysitosterol 7-Hydroxysitosterol 5,6-Epoxysitosterol 5,6␣-Epoxysitosterol 5␣,6-Dihydroxysitosterol 7-Ketositosterol a b c
Linear analysisa Calibration curve
R2
– y = (2.9x − 0.4) × 105 y = (1.7x + 0.9) × 105 y = (1.7x − 1.3) × 105 y = (1.6x + 0.7) × 105 y = (3.1x + 0.6) × 105 y = (2.6x − 1.0) × 105
– 0.9966 0.9984 0.9927 0.9986 0.9997 0.9954
Limit of detection (g/mL)
% Recovery of SPE purificationb
– 0.5 0.5 3 1 1 0.5
90.1 97.5 99.2 107.1 90.6 88.7 90.5
± ± ± ± ± ± ±
6.9 3.3 8.4 10.9 7.8 7.0 6.5
% Method recoveryc
87.2 92.5 90.2 115.3 102.3 85.5 87.2
± ± ± ± ± ± ±
3.0 5.0 8.3 14.8 6.7 5.3 4.0
Calibration curves were obtained by peak areas (y) vs. plotting concentration (x) (n = 10). Mean ± standard derivation (n = 10). Mean ± standard derivation (n = 7).
dihydroxy, and 7-ketositosterol). However, for 5,6epoxy, 5,6␣-epoxysitosterol, 19-hydroxycholesterol, and cholestanol, the selected quantification ions were not the base peaks (m/z 73 or 215) because these two ions were not characteristic. Their detection was then not selective enough. Other characteristic ions (m/z 413, 395, 353, and 356) were chosen for their quantitative analysis. The results showed that their RRF values were significantly different. They ranged from 0.12 to 6.44 with cholestanol or from 0.16 to 8.29 with 19-hydroxycholesterol in SIM mode. Even in TIC mode, their RRF values were also different from 0.62 to 1.99 with cholestanol and from 0.86 to 2.73 with 19-hydroxycholesterol. The response factors of 5,6␣-epoxy and 5,6-epoxysitosterol were astonishingly different. Then, the use of an arbitrary RRF equal to 1 within MS detection (SIM mode) will necessarily lead to important analytical errors. The RRF values were of utmost importance and taken into account for quantification of -sitosterol oxides in the following analysis. 3.2. Linearity, limit of detection, and recovery 19-Hydroxycholesterol is a good choice as an internal standard because it has quite the same analytical behaviour as cholesterol oxides [20,27,35,36]. Thus, it could be added directly to the food samples and then withstand similar saponification and purification processes. Moreover, its retention time was close to those of phytosterol oxides but without being co-eluted with them in the GC chromatogram. Table 3 summarizes the linearity and limit of detection for each -sitosterol oxide in the present study. The calibration of peak areas versus concentrations generated linear functions (coefficient of correlation r2 = 0.9927–0.9997) for all studied oxides within a range from limit of detection to 80 g/mL. The limits of detection (LOD) of the six oxides as their TMS ethers were estimated to be 0.5 g/mL for 7␣-hydroxy, 7-hydroxy, and 7-ketositosterol, 1 g/mL for 5,6␣-epoxy and 5␣,6-dihydroxy sitosterol, and 3 g/mL for 5,6hydroxysitosterol. A signal-to-noise ratio equal to 3 was used. As also shown in Table 3, most of the oxides had a relative recovery >90% in the SPE purification step. Average
method recoveries between 85 and 115% for each oxide when spiked in butter and assayed in complete analytical method with R.S.D. values below 10% (n = 7) in most cases indicated good recovery and repeatability of the method. In the quantitative calculation, the method recovery values were taken into account. 3.3. Optimization of the analytical method Generally, methods for determination of sterol oxides consist of saponification, lipid extraction, purification, identification by GC–MS, and quantification. In order to minimize the formation of artifacts and avoid the decomposition of the keto and epoxy derivatives during the saponification step [34,37,38], the saponification of lipids from oils was performed at room temperature overnight. The possibility of artifact formation was studied during this cold saponification and following the workup processes. Two hundred milligrams of butter were spiked with 1 mg pure -sitosterol and the analytical method was applied. Butter was selected as a research medium because it contains no phytosterols and consequently, no phytosterol oxides. GC results showed that no detectable level of -sitosterol oxides was present in the extracts. Moreover, the possibility of interconversion of each oxide during saponification and SPE purification was also evaluated. While each individual 20 g of -sitosterol oxide was separately spiked into 200 mg of butter and followed workup by the complete analytical method, no transformation happened between the oxides. Only the added oxide could be recovered in the GC analysis. Due to the analytes complexity in the food matrix, a cleanup step prior to GC analysis was mandatory. SiOH-SPE cartridges were used to isolate and concentrate polar lipids (containing phytosterol oxides) from low-polarity lipids and neutral lipid fraction (containing non-oxidized sterols) present in the unsaponifiable matter. In the work of Lampi et al. [20], an elution volume of 5 mL of hexane/diethyl ether (1:1, v/v) was used to remove the non-oxidized sterols from the cartridge prior to the oxide fraction. However, in the present study, this elution volume was reduced to 4 mL of cyclohexane/diethyl ether because trace amounts of 5,6␣epoxy and 5,6-epoxysitosterol were detected in the fifth
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milliliter of this elution. Ten replicate experiments and GC results showed no sterol oxides in the 4 mL of the cyclohexane/diethyl ether fraction (1:1, v/v), while all oxides were quantitatively removed from the SPE cartridge by the following 5 mL of acetone. 3.4. Formation of β-sitosterol oxides by heating β-sitosterol Phytosterols are basically stable compounds. However, under specific conditions, such as high temperature (>100 ◦ C) in the presence of air, oxidation process of phytosterols may occur in the same way as for cholesterol [2,24,38,39]. Some authors also reported the effect of refining and storage on the ppm level of the sterol oxides formed [31,33,40]. In order to study the effect of high temperatures on the formation of sterol oxides, 1 mg of pure -sitosterol was heated to 100, 150, or 200 ◦ C. No detectable level of oxides was found at 100 ◦ C in 30 min. At 150 ◦ C and 200 ◦ C, the main -sitosterol oxides represented during heating were 7␣-hydroxy, 7-hydroxy, 5,6␣-epoxy, 5,6-epoxy, and 7ketositosterol. No 5␣,6-dihydroxysitosterol was formed certainly due to the lack of water in the system. The major oxide formed at the two temperatures was 7-ketositosterol. The formation rate was quite constant during the first 30 min of heating at 150 ◦ C (Fig. 2(a)). In contrast, when -sitosterol was heated at 200 ◦ C, the oxide content increased very quickly during the first 15 min, then slightly decreased after 20–25 min. This phenomenon was probably due to thermodegradation and evaporation of the oxides formed (Fig. 2(b)). Similar results were shown in previous reports for the thermooxidation of stigmasterol or cholesterol [20,39]. 3.5. Quantification of β-sitosterol oxides in oils Vegetable oils or plant sterol-enriched foods are complex media because they contain several different phytosterols. -Sitosterol is among the most representative phytosterols, along with campesterol, stigmasterol, and other various types of minor phytosterols. The initial content of -sitosterol in the oils used was quantified by applying the analytical method to 200 mg of oil samples, and analyzing the fraction eluted with cyclohexane/diethyl ether (1:1, v/v) after spiking with 20 g of cholestanol as an internal standard. The total amounts of phytosterols were 2.6 and 1.6 mg/g in sunflower and olive oils, and the concentrations of -sitosterol were 1.5 and 0.9 mg/g in the oils, respectively. As mentioned above, the phytosterols contained in vegetable oils are labile through thermolytic and oxidation reactions when subjected to high temperatures or frying conditions. Fig. 3 presents the typical SIM chromatograms of the polar fractions in thermo-oxidized oils. The peaks of oxides were identified by their retention times as well as mass spectra, and quantitated with their characteristic ions of SIM mode along with their RRF, respectively, as shown in Tables 1 and 2.
Fig. 2. Major oxides formed by heating -sitosterol at different temperatures. Each -sitosterol oxide was quantified by capillary GC–MS with SIM mode: (a) 150 ◦ C, (b) 200 ◦ C. () 7␣-hydroxysitosterol; () 7hydroxysitosterol; () 5,6-epoxysitosterol; (×) 5,6␣-epoxysitosterol; () 7-ketositosterol.
Tables 4 and 5 summarize the concentration of six sitosterol oxides formed before and after heating of sunflower and olive oils at 150 and 200 ◦ C for 60 min. Initially, 7␣-hydroxy (6.5 g/g of oil), 7-hydroxy (7.0 g/g of oil), 7-keto (35.2 g/g of oil), and traces of 5␣,6dihydroxysitosterol (2.1 g/g of oil) were found in the native sunflower oil analyzed before heating. These oxides may be formed during oil processing and refinement [33,40]. No detectable levels of sterol oxides were found in the original olive oil. With heating, the amount of oxides formed in sunflower oil increased slowly during the first 30 min at 150 ◦ C and the first 15 min at 200 ◦ C. The total content was about 75 g/g of oil. After these durations, the content of total and individual -sitosterol oxides increased quickly at both temperatures with a total level about three times higher at 200 ◦ C (815.1 g/g of oil) than that at 150 ◦ C (240.9 g/g of oil) after 60 min. The content of 5␣,6-dihydroxysitosterol (2–10 g/g) did not change significantly at two temperatures. In olive oil, minor amounts of the oxides were formed after heating the olive oil at 150 ◦ C for 30 min or at 200 ◦ C for
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Fig. 3. Typical selection ion monitoring (SIM) chromatograms of m/z 356 + 353 + 485 + 413 + 395 + 432 + 396 obtained by GC–MS analysis of (a) six sitosterol oxides and internal standards (20 g of each compound), as well as in (b) the polar fractions of the heated sunflower oil, and in (c) the polar fraction of the heated olive oil. The upper chromatograms are corresponding total ion chromatogram (TIC) of oxides standards and polar fractions in heated oils. Chromatographic peaks: (1) cholestanol (internal standard); (2) 19-hydroxycholesterol (internal standard); (3) 7␣-hydroxysitosterol; (4) 7-hydroxysitosterol; (5) 5,6-epoxysitosterol; (6) 5,6␣-epoxysitosterol; (7) 5␣,6-dihydroxysitosterol; (8) 7-ketositosterol.
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Table 4 Concentrations of -sitosterol oxides (g/g of oil) after heating sunflower oil at 150 or 200 ◦ C for different durationsa Analytes
150 ◦ C
200 ◦ C
0 min
5 min
15 min
30 min
60 min
0 min
5 min
15 min
30 min
60 min
7␣-Hydroxysitosterol 7-Hydroxysitosterol 5,6-Epoxysitosterol 5,6␣-Epoxysitosterol 5␣,6-Dihydroxysitosterol 7-Ketositosterol
6.5 ± 0.3 7.0 ± 0.3 n.d. n.d. 2.1 ± 0.4 35.2 ± 2.4
9.7 ± 0.6 9.6 ± 0.7 2.1 ± 0.2 1.2 ± 0.0 1.8 ± 0.1 41.7 ± 1.3
9.8 ± 0.2 9.4 ± 0.3 2.9 ± 0.2 2.0 ± 0.2 1.8 ± 0.0 44.0 ± 1.1
10.3 ± 0.4 10.2 ± 0.5 3.1 ± 0.6 1.1 ± 0.1 2.0 ± 0.2 45.9 ± 2.2
31.6 ± 2.3 49.5 ± 5.0 45.0 ± 4.3 24.3 ± 4.8 3.9 ± 0.6 86.6 ± 8.3
6.5 ± 0.3 7.0 ± 0.3 n.d. n.d. 2.1 ± 0.4 35.2 ± 2.4
10.1 ± 0.4 9.7 ± 0.5 2.9 ± 0.2 1.1 ± 0.0 1.8 ± 0.0 41.8 ± 1.0
10.6 ± 0.3 11.5 ± 0.4 2.2 ± 0.2 1.2 ± 0.1 2.1 ± 0.2 49.4 ± 0.7
33.1 ± 3.7 46.2 ± 3.8 46.9 ± 2.3 20.2 ± 2.2 3.8 ± 0.4 72.9 ± 1.4
81.3 ± 9.3 98.9 ± 8.7 220.3 ± 11.2 147.3 ± 21.5 9.1 ± 0.8 258.2 ± 16.5
Total amount (g/g oil) % -sitosterol oxidized
50.8 2.0
66.1 2.3
69.9 2.4
72.6 2.6
240.9 9.5
50.8 2.0
67.4 2.4
77.0 2.7
223.1 7.6
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n.d. = not detected. a Mean ± standard derivation (n = 3).
Table 5 Concentration of -sitosterol oxides (g/g oil) after heating olive oil at 150 or 200 ◦ C for different durationsa Analytes
150 ◦ C
200 ◦ C
0 min
5 min
15 min
30 min
60 min
0 min
5 min
15 min
30 min
60 min
7␣-Hydroxysitosterol 7-Hydroxysitosterol 5,6-Epoxysitosterol 5,6␣-Epoxysitosterol 5␣,6-Dihydroxysitosterol 7-Ketositosterol
n.d. n.d. n.d. n.d. n.d. n.d.
0.5 ± 0.0 0.5 ± 0.0 n.d. n.d. n.d. 0.7 ± 0.1
0.6 ± 0.1 0.6 ± 0.1 n.d. n.d. n.d. 1.0 ± 0.0
0.9 ± 0.1 1.1 ± 0.2 1.5 ± 0.3 1.0 ± 0.1 n.d. 1.5 ± 0.2
5.4 ± 0.2 9.5 ± 0.2 8.0 ± 0.4 3.8 ± 0.3 0.4 ± 0.3 10.0 ± 0.3
n.d. n.d. n.d. n.d. n.d. n.d.
0.5 ± 0.1 0.6 ± 0.2 n.d. n.d. n.d. 0.7 ± 0.2
0.8 ± 0.1 1.1 ± 0.2 0.6 ± 0.2 0.2 ± 0.3 n.d. 1.4 ± 0.2
21.1 ± 2.4 32.6 ± 3.0 31.5 ± 1.8 17.7 ± 0.2 2.3 ± 0.0 20.5 ± 1.0
50.3 ± 1.1 54.8 ± 1.2 94.5 ± 5.7 56.6 ± 5.6 3.2 ± 0.5 105.9 ± 9.3
Total amount (g/g oil) % -sitosterol oxidized
n.d. 0
1.7 1.0
2.2 1.3
6.0 3.0
37.1 7.2
n.d. 0
1.8 1.1
4.1 2.4
125.7 4.8
365.3 20.3
n.d. = not detected. a Mean ± standard derivation (n = 3).
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15 min, but they contributed to less than 3% of the -sitosterol oxidized. Conversely, the sunflower oil already contained similar levels of these oxides in its initial state. Trace levels of 5␣,6-dihydroxysitosterol were found in olive oil (<4 g/g) after heating. The result also showed that higher temperature will lead to more oxides formed. The total amounts of -sitosterol oxides in olive oil were 37.1 and 365.3 g/g of oil for heating at 150 and 200 ◦ C after 60 min, respectively. In the present heating assay of -sitosterol and oils, the  configuration of the oxides formed more favorably than that of the corresponding ␣ oxides. This result was in agreement with the previous report that formation of the -epimer of cholesterol epoxide was favored over the ␣-epimer [6]. The main oxides formed were the same at 150 and 200 ◦ C in the two oils. At 150 ◦ C, 7-hydroxy and 7-ketositosterol were present in the highest concentrations, while at 200 ◦ C, 7-ketositosterol quantitatively predominated, followed by 5,6-epoxysitosterol. As the effect of cooking time on the oxidation of cholesterol in meats [36], the formation rate of phytosterol oxides in plant oils differed according to the heating time and temperatures. Lower temperature and shorter periods might avoid the rapid formation of phytosterol oxides. However, significant amounts of oxides were detected in the oils heated at 200 ◦ C for 60 min (20–25% of -sitosterol oxidized). The total content of -sitosterol oxides formed at 200 ◦ C for 60 min with sunflower oil was about two times higher than that of olive oil under the same conditions.
Acknowledgements The authors thank Dr. Jennifer Wytko for her help in the preparation of the manuscript. This work was supported by a grant from the Minist`ere de la Jeunesse, de l’Education Nationale et de la Recherche, France (RARE 015 No. 02 P 0640).
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[9] Guardiola F, Dutta PC, Savage GP, Codony R, editors. Cholesterol and phytosterol oxidation products in foods and biological samples: analysis, occurrence and biological effects. Champaign, IL: AOCS Press; 2002. [10] Guardiola F, Codony R, Addis PB, Rafecas M, Boatell J. Biological effects of oxysterols: current status. Food Chem Toxicol 1996;34:193–211. [11] Brown AJ, Jessup W. Oxysterols and atherosclerosis. Atherosclerosis 1999;142:1–28. [12] Ling WH, Jones PJH. Dietary phytosterols: a review of metabolism, benefits and side effects. Life Sci 1995;57:195–206. [13] Piironen V, Lindsay DG, Miettinen TA, Toivo J, Lampi AM. Plant sterols: biosynthesis, biological function and their importance to human nutrition. J Sci Food Agric 2000;80:939–66. [14] Moreau RA, Whitaker BD, Hicks KB. Phytosterol, phytostanols, and their conjugates in food: structural diversity, quantitative analysis, and health-promoting uses. Prog Lipid Res 2002;41:457–500. [15] Adcox C, Boyd L, Oehrl L, Allen J, Fenner G. Comparative effects of phytosterol oxides and cholesterol oxides in cultured macrophagederived cell lines. J Agric Food Chem 2001;49:2090–5. [16] Grandgirard A, Martine L, Demaison L. Oxyphytosterols are present in plasma of healthy human subjects. Br J Nutr 2004;91:101–6. [17] Shan H, Pang J, Li S, Chiang TB, Wilson WK, Schroepfer Jr GJ. Chromatographic behaviour of oxygenated derivatives of cholesterol. Steroids 2003;68:221–33. [18] Dutta PC. Studies on phytosterol oxides. II. Content in some vegetable oils and in French fries prepared in these oils. J Am Oil Chem Soc 1997;74:659–66. [19] Plat J, Brzezinka H, L¨utjohann D, Mensink RP, Bergmann K. Oxidized plant sterols in human serum and lipid infusions as measured by combined gas-liquid chromatography–mass spectrometry. J Lipid Res 2001;42:2030–8. [20] Lampi AM, Juntunen L, Toivo J, Piironen V. Determination of thermo-oxidation products of plant sterols. J Chromatogr B 2002;777:83–92. [21] Johnsson L, Dutta PC. Characterisation of side-chain oxidation products of sitosterol and campesterol by chromatographic and spectroscopic methods. J Am Oil Chem Soc 2003;80:767–76. [22] Grandgirard A, Martine L, Joffre C, Juaneda P, Berdeaux O. Gas chromatographic separation and mass spectrometric identification of mixtures of oxyphytosterols and oxycholesterol derivatives. Application to a phytosterol-enriched food. J Chromatogr A 2004;1040:239–50. [23] Johannes C, Lorenz RL. Preparation and mass spectrometry of 14 pure and 18 O2 -labeled oxidation products from the phytosterols sitosterol and stigmasterol. Anal Biochem 2004;325:107–16. [24] Johnsson L, Dutta PC. Separation of phytosterol oxidation products by combination of different polarity gas chromatography capillary columns. J Chromatogr A 2005;1064:213–7. [25] Guardiola F, Bou R, Boatella J, Codony R. Analysis of sterol oxidation products in foods. J AOAC Int 2004;87:441–66. [26] Diczfalusy U. Analysis of cholesterol oxidation products in biological samples. J AOAC Int 2004;87:467–73. [27] Regueiro JAG, Maraschiello C. Procedure for the determination of eight cholesterol oxides in poultry meat using on-column and solvent venting capillary gas chromatography. J Chromatogr A 1997;764:279–93. [28] Turchetto E, Lercker G, Bortolomeazzi R. Oxysterol determination in selected coffees. Toxicol Ind Health 1993;9:519–27. [29] Zhang X, Geoffroy P, Miesch M, Julien-David D, Raul F, Aoud´eWerner D, et al. Gram-scale chromatographic purification of sitosterol. Synthesis and characterization of -sitosterol oxides. Steroids 2005;70:886–95. [30] Bortolomeazzi R, Zan MD, Pizzale L, Conte LS. Mass spectrometry characterization of the 5␣, 7␣, and 7-hydroxy derivatives of sitosterol, campesterol, stigmasterol and brassicasterol. J Agric Food Chem 1999;47:3069–74.
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Identification and quantitative analysis of -sitosterol oxides in vegetable oils by capillary gas chromatography–mass spectrometry Xin Zhang a , Diane Julien-David a , Michel Miesch b , Philippe Geoffroy b , Francis Raul c , Stamatiki Roussi c , Dalal Aoud´e-Werner d , Eric Marchioni a,∗ a
b
Laboratoire de Chimie Analytique et Sciences de l’Aliment (UMR 7512), Facult´e de Pharmacie, Universit´e Louis Pasteur, 74 route du Rhin, 67400 Illkirch, France Laboratoire de Chimie Organique Synth´etique (UMR 7123), Universit´e Louis Pasteur, 1 rue Blaise Pascal, 67008 Strasbourg, France c Laboratoire d’Oncologie Nutritionnelle, Inserm UMR S392/IRCAD, 1 place de l’Hˆ opital, 67091 Strasbourg, France d A´ erial, rue Laurent Fries—Parc d’Innovation, 67412 Illkirch, France Received 28 October 2004; received in revised form 9 June 2005; accepted 9 June 2005 Available online 20 July 2005
Abstract As vegetable oils and phytosterol-enriched spreads are marketed for frying food or cooking purposes, temperature is one of the most important factors leading to the formation of phytosterol oxides in food matrix. A methodology based on saponification, organic solvent extraction, solid-phase extraction (SPE), followed by mass spectrometric identification and quantitation of -sitosterol oxides using capillary gas chromatography–mass spectrometry (GC–MS) in selected ion monitoring (SIM) mode was developed and characterized. Relative response factors of six -sitosterol oxides, including 7␣-hydroxy, 7-hydroxy, 5,6␣-epoxy, 5,6-epoxy, 7-keto, and 5␣,6-dihydroxysitosterol, were calculated against authentic standards of 19-hydroxycholesterol or cholestanol. Linear calibration data, limit of detection, and sample recoveries during analytical process. Recoveries of these oxidation compounds in spiked samples ranged from 88 to 115%, while relative standard derivation (R.S.D.) values were below 10% in most cases. The analytical method was applied to quantify -sitosterol oxides formed in thermal-oxidized vegetable oils which were heated at different temperatures and for varying time periods. Sitosterol oxidation is strikingly higher in sunflower oil relative to olive oil under all conditions of temperature and heating time. © 2005 Elsevier Inc. All rights reserved. Keywords: -Sitosterol oxides; Gas chromatography–mass spectrometry (GC–MS); Selected ion monitoring (SIM) mode; Quantification; Oil; Thermaloxidation
1. Introduction Phytosterols in edible vegetable oils or plant sterolenriched foods are structurally similar to cholesterol, the only differences being the position of double bond and the alkyl side chain. Both cholesterol and phytosterols contain an unsaturated ring-structure and are thus susceptible undergo oxidation in the presence of oxygen. Oxidation is accelerated by heating, exposure to ionizing radiation, light, chemical catalysts, or enzymatic processes [1–5]. For cholesterol, more than 80 oxides have been identified and extensively ∗
Corresponding author. Tel.: +33 3 90244326; fax: +33 3 90244325. E-mail address: [email protected] (E. Marchioni).
0039-128X/$ – see front matter © 2005 Elsevier Inc. All rights reserved. doi:10.1016/j.steroids.2005.06.004
studied and reviewed [6–9]. The main oxides are hydroxy, keto, and epoxy compounds. The latter may hydrate to dihydroxy sterols [6]. The in vitro studies have shown that some cholesterol oxides have cytotoxic, mutagenic, atherogenic, and carcinogenic activities [10,11]. Due to the structural similarity with cholesterol and its oxides, phytosterols and their oxides have received much attention during the last decades with respect to their biological properties and consequently, their performance in food safety evaluation [12–16]. Some studies [9,12] have shown that supplementation of foods with phytosterols and phytosterol esters may induce the presence of their oxides in foods and human plasma. The analytical methods for phytosterol oxides are based on those developed for cholesterol oxides. Gas chromatography
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(GC) or high performance liquid chromatography (HPLC) is most often employed for the separation and quantification of cholesterol oxides. Although HPLC provides good resolution for cholesterol oxides [1,17], this technique is not suitable for phytosterol oxides analysis because of excessive interference, modest sensitivity, and limited chromatographic resolution. For this reason, final separation and identification of phytosterol oxides in food matrix are always carried out by GC–FID or GC–MS [18–24]. Guardiola et al. [25] and Diczfalusy [26] reviewed the analysis of sterol oxides in foods and biological samples. Studies dealing with sterol oxide mixtures from complex food samples using GC–MS operating in selected ion monitoring (SIM) mode as quantitative technique may increase the detection selectivity and avoid the numerous interfering problems [27,28]. Since individual standards of phytosterol oxides are not available commercially, most research works, including chemical quantitative analysis, biochemical, and medical studies, have used mixtures of phytosterol oxides [15,16,20]. Six individual -sitosterol oxides have been synthesized recently [29]. In the present study, these compounds were used as standards, a precise quantitative analytical method of these products formed in thermal-oxidized oils are performed. Much attention was paid to the formation of oxides when heating vegetable oils for short periods of time (≤1 h).
2. Experimental 2.1. Materials and reagents 5␣-Cholestan-3-ol (cholestanol, 95%) and cholest-5ene-3,19-diol (19-hydroxycholesterol, 95%), purchased from Sigma (Steinheim, Germany), were used as internal standards. -Sitosterol was purified from Generol 95R (Cognis, Saint-Fargeau-Ponthierry, France), a commercial phytosterol mixture, in the laboratory [29]. -Sitosterol oxides, e.g., sitost-5-ene-3,7␣-diol (7␣-hydroxysitosterol), sitost-5-ene-3,7-diol (7-hydroxysitosterol), 5,6␣-epoxy5␣-sitostan-3-ol (5,6␣-epoxysitosterol), 5,6-epoxy-5sitostan-3-ol (5,6-epoxysitosterol), 3-hydroxysitost-5en-7-one (7-ketositosterol), and sitost-5-en-3,5␣,6-triol (5␣,6-dihydroxysitosterol), were then chemically synthesized [29]. The purity of each compound was ∼95% as determined by 1 H NMR, 13 C NMR, and GC–MS. Stock solution (1 mg/mL) of each analyte and internal standards, cholestanol and 19-hydroxycholesterol, were prepared separately in ethyl acetate and stored at 4 ◦ C in the dark. Ethyl acetate was obtained from Acros (Geel, Belgium), ethanol from Carlo Erba (France), cyclohexane from Fluka (Buchs, Switzerland), diethyl ether from Riedel-de Ha¨en (Seelze, Germany), acetone and pyridine from Prolabo (Fontenay sous Bois, France), isooctane and potassium hydroxide (KOH) from Merck (Darmstadt, Germany), and anhydrous sodium sulfate (Na2 SO4 ) from SDS (Peypin, France). All organic solvents and reagents were
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of analytical grade. Ultra-pure water was obtained by means of a Millipore Q water purification system (Milford, MA, USA). N-Methyl-N-(trimethylsilyl)trifluoroacetamide (MSTFA), used as the silylation reagent, was provided by Sigma-Aldrich (Steinheim, Germany). The SiOH-SPE cartridges (3 mL/500 mg) were purchased from MachereyNagel (Chromabond® , D¨uren, Germany). 2.2. Food samples Sunflower oil (1 L, Lesieur Tournesol, France), olive oil (0.5 L, Puget, France), and butter (250 g, Pr´esident, France) were of edible quality and purchased in a local supermarket. 2.3. Samples pretreatment Firstly, 1 mg of -sitosterol was weighed in an open glassware (10 mm × 60 mm i.d.) and heated in an oven (Memmert, Schwabach, Germany) which had been set at 100, 150, or 200 ◦ C for 5, 15, and 30 min. The final products were converted to TMS ethers (as described below) and analyzed directly by GC–MS to estimate the main oxides formed. After the model assay, approximately 200 mg of oil samples (sunflower oil or olive oil) was weighted in the same type of glassware to ensure efficient surface contact with air, and then, they were placed into the oven and heated at 150 or 200 ◦ C. After various durations (from 5 to 60 min), the glasswares were removed from the oven, quickly covered and sealed with aluminium foil, and allowed to cool to room temperature. The samples were then dissolved in excess ethyl acetate, collected into 30 mL brown vials, evaporated to dryness under a gentle nitrogen flow, and stored at 4 ◦ C under nitrogen. Each sample was assayed in triplicate. 2.4. Isolation and purification of phytosterol oxides from food samples The procedure for sample preparation described here for phytosterol oxides analysis was adapted from the methods proposed by Dutta [18] and Lampi et al. [20] with minor changes. The detailed processes were as follows: • Cold saponification. Prior to saponification, 20 L of 19hydroxycholesterol solution (1 mg/mL in ethyl acetate) as an internal standard was spiked into the heated 200 mg of oil samples in a 30 mL brown vial. After removal of the ethyl acetate under nitrogen, 9 mL of ethanol, and 0.5 mL of saturated KOH aqueous solution were added into the vial. The sample was purged with nitrogen, and the tightly closed brown vial was put into a rotary shaker (Edmund B¨uhler, Johanna Otta GmbH, Hechingen, Germany) at ambient temperature in the dark overnight (15 h). • Extraction. The solution was transferred in a 100 mL separation funnel and 10 mL of distilled water were added. The unsaponifiable fraction was extracted with diethyl ether (4× 15 mL). The combined organic extracts were
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washed with 20 mL of 0.5 M aqueous KOH solution and 0.2 M aqueous Na2 SO4 solution (3× 15 mL). The organic solvents were removed with a rotary vacuum evaporator (25 ◦ C, 200 mbar), and the residue was dried under nitrogen flow. After additional 2 mL of diethyl ether, the unsaponifiable residue was carefully transferred using a pipette to a 2 mL glass test tube and then evaporated to dry matter under nitrogen flow. • Solid-phase extraction (SPE) purification. The SiOH-SPE cartridges were first conditioned with 5 mL of cyclohexane. The unsaponifiable matter was dissolved in 1 mL of a solution of cyclohexane/diethyl ether (9:1, v/v) and was loaded onto the cartridge. Low-polarity lipids and non-oxidized sterols were eluted with 5 mL of cyclohexane/diethyl ether (9:1, v/v) followed by 4 mL of cyclohexane/diethyl ether (1:1, v/v). Finally, the sterol oxides were eluted out of the cartridge with 5 mL of acetone and collected into a test tube to which 20 L of cholestanol (1 mg/mL in ethyl acetate) were added. This residue was dried under a gentle nitrogen flow. • Derivatization. The purified sterol oxides residue was converted to trimethylsilyl (TMS) ethers with 50 L of pyridine and 40 L of MSTFA at room temperature in the dark overnight and then, diluted with 410 L of isooctane. One microliter of each sample was injected into the GC–MS system. 2.5. Gas chromatography–mass spectrometry (GC–MS) analysis GC–MS analyses were performed on a Varian STAR 3400 GC instrument equipped with an on-column SPI injector coupled to a Varian SATURN 2000 mass sensitive detector (Varian, France). Data acquisition and processing, and instrumental control were performed by Varian Saturn WS software. Analytes were separated in a VF-5ms capillary column (phase stationary: 5% phenyl–95% dimethylpolysiloxane, thickness of 0.1 m, 60 m × 0.25 mm, Varian, France). The column temperature gradient was programmed from 105 ◦ C (hold for 2 min) to 170 ◦ C at 20 ◦ C/min and then, to 320 ◦ C at 7 ◦ C/min (hold for 15 min). The injector operating conditions were as follows: injection volume 1 L; initial injector temperature of 105 ◦ C was increased to 300 ◦ C at 100 ◦ C/min (hold for 40 min). Helium (purity 99.9995%) was used as a carrier gas with a flow rate of 1 mL/min. Electronic impact (EI) ionization mode mass spectra were obtained at 70 eV and monitored on the full-scan range (m/z 40–600). Quantitative data were carried out using selected ion monitoring (SIM) analysis. The analytes were quantified as follows: Ac τis nc = nis RRFc Ais τc where nc and nis were the moles of component analyzed in the sample and of internal standard added to the sample, respectively; Ac and Ais the peak areas of component analyzed in
the sample and of internal standard added to the sample, respectively; τ c and τ is the method recoveries of component analyzed in the sample and of internal standard added to the sample, respectively; and RRFc was the relative response factor of each pure -sitosterol oxide compared to the internal standard, which was calculated as follows: RRFc =
nc Ais RRFis nis Ac
where Ac and Ais were the peak areas of standard oxide and of internal standard, respectively; nc and nis the moles of standard oxide and of internal standard, respectively; and RRFis was the response factor of the internal standard, which was set to 1. 2.6. Linearity, limit of detection and recovery For each oxide, standard solutions were made at six different concentrations between 0.1 and 100 g/mL, derivatized, and assayed in duplicate. Linear plots of peak areas versus concentration were calculated. Recoveries of SiOH-SPE purification were tested with mixtures containing the six sitosterol oxides as well as 19-hydroxycholesterol (10, 20, and 40 g of each one) spiked with 20 g of cholestanol as an internal standard. Recoveries of the complete analytical method were carried out in 200 mg of butter spiked with the six -sitosterol oxides (10, 20, and 40 g) and 20 g of 19hydroxycholesterol as an internal standard.
3. Results and discussion 3.1. Identification and characterization of β-sitosterol oxides by GC–MS A series of individual -sitosterol oxides (∼95% purity), including 7␣-hydroxy, 7-hydroxy, 5,6␣-epoxy, 5,6-epoxy, 7-keto, and 5␣,6-dihydroxysitosterol, were synthesized from purified -sitosterol and characterized by means of GC–MS as TMS ethers. Chromatographic separations of these compounds were well achieved on a VF-5ms capillary column and identified by their MS data. Retention times were consistent within 1.2 s across runs. The typical retention times and selected ions chosen for identification and quantitation along with their relative abundances are summarized in Table 1. The mass spectra of these oxides showed the similar typical fragmentation patterns, which were in agreement with the data reported previously [2,30,31] with minor differences in the relative abundances. One point should be mentioned that some mass spectra rounded numbers to the unit, giving occurrence to impaired mass fragments of molecular ion or base peak. For the mass spectra of 7␣-hydroxy and 7-hydroxysitosterol as di-TMS ether derivatives, the base peak was observed at m/z 485 (M+ − TMSOH). The relative intensity of molecular ions (M+ ) at m/z 575 was 1%.
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Table 1 Retention time (tR ) and relative retention time (RRT) of -sitosterol and its oxides as TMS ethers in relation to that of 19-hydroxycholesterol, as well as their specific ions Analytes
tR (min)
RRTa
Mw b
SIM ions (relative abundance, %) Identification
Quantitation
-Sitosterol Cholestanol 19-Hydroxycholesterol 7␣-Hydroxysitosterol 7-Hydroxysitosterol 5,6-Epoxysitosterol 5,6␣-Epoxysitosterol 5␣,6-Dihydroxysitosterol 7-Ketositosterol
31.42 29.28 29.55 30.48 32.03 33.08 33.32 34.07 35.58
1.06 0.99 1.00 1.03 1.08 1.11 1.12 1.15 1.20
486.7 460.7 546.7 574.7 574.7 502.7 502.7 664.7 500.7
382 (52), 397 (100), 472 (16), 487 (32) 215 (100), 356 (65), 446 (29), 461 (12) 73 (100), 353 (75), 367 (59), 547 (3) 470 (4), 485 (100), 575 (1) 470 (3), 485 (100), 575 (1) 73 (100), 395 (38), 413 (52), 503 (24) 73 (100), 395 (60), 413 (46), 503 (15) 432 (100), 485 (84), 560 (34), 575 (13) 396 (100), 486 (41), 501 (80)
397 356 353 485 485 413 395 432 396
a b
Relative to the retention time of 19-hydroxycholesterol TMS ether. Mw : molecular weight as TMS ether.
The other typical fragments 470 (M+ − TMSOH-CH3 ), 395 (M+ − 2TMSOH), 253 (M+ − 2TMSOH-side chain), and 129 (5-sterol) were of low intensity (1–6%), except for m/z 73 (22%). For the TMS ether derivatives of 5,6␣-epoxy and 5,6-epoxysitosterol, similar ion fragmentation patterns were found, apart from some differences in their relative peak intensities. The molecular ion at m/z 503 was present for both compounds (15 and 24%), and the base peak was at m/z 73. The molecular ion of 7-ketositosterol TMS ether was at m/z 501 (M+ , 80%), and the base peak was at m/z 396 (M+ − TMSOH-CH3 , 100%). This result was different from previously reported data [28,31] in which an unspecified base peak at m/z 174 was observed. The other typical fragments were consistent with the data reported previously [31,32]. Special attention should be taken with 5␣,6dihydroxysitosterol. Indeed, depending on the silylating reagents and on the reaction time used, bis- and tris-TMS ether derivatives may be produced from 5␣,6dihydroxycholesterol or 5␣,6-dihydroxysitosterol because the derivatization at the 5␣-hydroxyl group of the sterol is sterically hindered [31,32]. In the present work, when derivatization was performed at 50 ◦ C for 60 min, three
hydroxyl groups of 5␣,6-dihydroxysitosterol could not completely be derivatized to the tris-TMS ether derivative. A gas chromatogram showed two peaks, one (tR = 33.98 min) corresponding to tris-TMS ether (with three OTMS groups) and the other (tR = 37.21 min) to the bis-TMS ether (with one hydroxy and two OTMS groups) (Fig. 1(a)). To avoid this phenomenon, the derivation reaction between analytes and MSTFA was carried out at ambient temperature overnight in the dark. The three hydroxyl groups of 5␣,6-dihydroxysitosterol could then be converted to tris-TMS ether (Fig. 1(b)). Its base peak was observed at m/z 432 (M+ − TMSOH-side chain). The other major fragments as well as their relative intensities were at m/z 575 (M+ − TMSOH, 13%), 560 (M+ − TMSOH-CH3 , 34%), 485 (M+ − 2TMSOH, 84%), 470 (M+ − 2TMSOH-CH3 , 28%), 395 (M+ − 3TMSOH, 11%), and 380 (M+ − 3TMSOH-CH3 , 4%). Moreover, this method was suitable for the silylation of all other studied oxides present in the samples prior to GC analysis. Relative response factors (RRF) of six -sitosterol oxides were calculated with SIM mode in the present work in relation to the internal standard, cholestanol or 19hydroxycholesterol, are listed in Table 2.
Table 2 Relative response factors (RRF) of -sitosterol and its oxides as TMS ethers in relation to those of cholestanol or 19-hydroxycholesterola Analytes
-Sitosterol Cholestanol 19-Hydroxycholesterol 7␣-Hydroxysitosterol 7-Hydroxysitosterol 5,6-Epoxysitosterol 5,6␣-Epoxysitosterol 5␣,6-Dihydroxysitosterol 7-Ketositosterol a b c
RRF (SIM mode)c
RRF (TIC mode) Relative to cholestanol
Relative to 19hydroxycholesterol
Relative to cholestanol
Relative to 19hydroxycholesterol
1.06 ± 0.04b – 0.69 ± 0.05 0.79 ± 0.05 0.85 ± 0.07 1.99 ± 0.31 1.25 ± 0.09 0.62 ± 0.29 1.31 ± 0.09
– 1.45 ± 0.08 – 1.05 ± 0.09 1.18 ± 0.08 2.73 ± 0.17 1.66 ± 0.14 0.86 ± 0.07 1.75 ± 0.13
0.17 ± 0.01b – 0.77 ± 0.03 0.12 ± 0.01 0.13 ± 0.01 6.44 ± 0.20 3.54 ± 0.08 0.35 ± 0.02 1.00 ± 0.08
– 1.29 ± 0.05 – 0.16 ± 0.01 0.17 ± 0.01 8.29 ± 0.28 4.63 ± 0.16 0.49 ± 0.02 1.31 ± 0.12
Mean ± standard derivation (n = 10). Mean ± standard derivation (n = 3). The characteristic ions for quantitation in SIM mode, see Table 1.
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Fig. 1. Influence of silylation conditions on the derivatization of 5␣,6-dihydroxysitosterol: (a) 50 ◦ C for 60 min, (b) room temperature for overnight. The first silylation condition lead to the formation of two peaks as shown in GC chromatogram (a). The upper left mass spectrum corresponds to tris-TMS ether (peak 1, with three OTMS groups). The upper right mass spectrum corresponds to bis-TMS ether (peak 2, with one hydroxy and two OTMS groups).
In previous works, because the phytosterol oxides were not commercially available, the RRF values of these compounds were usually adopted from those of the corresponding cholesterol oxides [19,20] or assumed to be a value of 1 [33,34] in GC–FID or GC–MS analysis with total ion chromatogram (TIC) mode. However, although good separation and analytical methods had been reported for cholesterol oxides [6,7,9], the chromatographic resolution problems associated with the analysis of phytosterol oxides in foods of plant origin or mixed origin (animal/plant-based foods) are quite complex because of the high probabilities of co-elution [9,22]. Such as, in the present GC separation system, 5,6-epoxysitosterol was co-eluted with 7-hydroxycampesterol. Therefore, due to high relative intensity of some characteristic ions, SIM mode has advantages in sensitivity, identification and quantitation of sterol oxides in food matrix. Guardiola et al. [25]
reviewed the characteristic ions (m/z) used to quantify cholesterol oxides as trimethylsilyl ether derivatives by GC–MS. However, in relation to phytosterol oxides, the literature is scarce. The only work that has quantified 7-ketositosterol in food samples by GC–MS operated in SIM mode was performed by Turchetto et al. [28], who selected the characteristic ions at m/z 500, 484, 410, and 395 to quantify the TMSE derivative of this compound. Since six -sitosterol oxides were synthesized and characterized recently, their exact RRF values were available. The ions selected for quantification were m/z 353 for 19-hydroxycholesterol, m/z 485 for 7␣hydroxy and 7-hydroxy, m/z 413 for 5,6-epoxy, m/z 395 for 5,6␣-epoxy, m/z 432 for 5␣,6-dihydroxy, and m/z 396 for 7-ketositosterol. In some cases, their base peaks were chosen (7␣-hydroxy, 7-hydroxy, 5␣,6-
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Table 3 Analytical parameters of gas chromatography for each -sitosterol oxide as TMS ethers assayed by the present method Analytes
19-Hydroxycholesterol 7␣-Hydroxysitosterol 7-Hydroxysitosterol 5,6-Epoxysitosterol 5,6␣-Epoxysitosterol 5␣,6-Dihydroxysitosterol 7-Ketositosterol a b c
Linear analysisa Calibration curve
R2
– y = (2.9x − 0.4) × 105 y = (1.7x + 0.9) × 105 y = (1.7x − 1.3) × 105 y = (1.6x + 0.7) × 105 y = (3.1x + 0.6) × 105 y = (2.6x − 1.0) × 105
– 0.9966 0.9984 0.9927 0.9986 0.9997 0.9954
Limit of detection (g/mL)
% Recovery of SPE purificationb
– 0.5 0.5 3 1 1 0.5
90.1 97.5 99.2 107.1 90.6 88.7 90.5
± ± ± ± ± ± ±
6.9 3.3 8.4 10.9 7.8 7.0 6.5
% Method recoveryc
87.2 92.5 90.2 115.3 102.3 85.5 87.2
± ± ± ± ± ± ±
3.0 5.0 8.3 14.8 6.7 5.3 4.0
Calibration curves were obtained by peak areas (y) vs. plotting concentration (x) (n = 10). Mean ± standard derivation (n = 10). Mean ± standard derivation (n = 7).
dihydroxy, and 7-ketositosterol). However, for 5,6epoxy, 5,6␣-epoxysitosterol, 19-hydroxycholesterol, and cholestanol, the selected quantification ions were not the base peaks (m/z 73 or 215) because these two ions were not characteristic. Their detection was then not selective enough. Other characteristic ions (m/z 413, 395, 353, and 356) were chosen for their quantitative analysis. The results showed that their RRF values were significantly different. They ranged from 0.12 to 6.44 with cholestanol or from 0.16 to 8.29 with 19-hydroxycholesterol in SIM mode. Even in TIC mode, their RRF values were also different from 0.62 to 1.99 with cholestanol and from 0.86 to 2.73 with 19-hydroxycholesterol. The response factors of 5,6␣-epoxy and 5,6-epoxysitosterol were astonishingly different. Then, the use of an arbitrary RRF equal to 1 within MS detection (SIM mode) will necessarily lead to important analytical errors. The RRF values were of utmost importance and taken into account for quantification of -sitosterol oxides in the following analysis. 3.2. Linearity, limit of detection, and recovery 19-Hydroxycholesterol is a good choice as an internal standard because it has quite the same analytical behaviour as cholesterol oxides [20,27,35,36]. Thus, it could be added directly to the food samples and then withstand similar saponification and purification processes. Moreover, its retention time was close to those of phytosterol oxides but without being co-eluted with them in the GC chromatogram. Table 3 summarizes the linearity and limit of detection for each -sitosterol oxide in the present study. The calibration of peak areas versus concentrations generated linear functions (coefficient of correlation r2 = 0.9927–0.9997) for all studied oxides within a range from limit of detection to 80 g/mL. The limits of detection (LOD) of the six oxides as their TMS ethers were estimated to be 0.5 g/mL for 7␣-hydroxy, 7-hydroxy, and 7-ketositosterol, 1 g/mL for 5,6␣-epoxy and 5␣,6-dihydroxy sitosterol, and 3 g/mL for 5,6hydroxysitosterol. A signal-to-noise ratio equal to 3 was used. As also shown in Table 3, most of the oxides had a relative recovery >90% in the SPE purification step. Average
method recoveries between 85 and 115% for each oxide when spiked in butter and assayed in complete analytical method with R.S.D. values below 10% (n = 7) in most cases indicated good recovery and repeatability of the method. In the quantitative calculation, the method recovery values were taken into account. 3.3. Optimization of the analytical method Generally, methods for determination of sterol oxides consist of saponification, lipid extraction, purification, identification by GC–MS, and quantification. In order to minimize the formation of artifacts and avoid the decomposition of the keto and epoxy derivatives during the saponification step [34,37,38], the saponification of lipids from oils was performed at room temperature overnight. The possibility of artifact formation was studied during this cold saponification and following the workup processes. Two hundred milligrams of butter were spiked with 1 mg pure -sitosterol and the analytical method was applied. Butter was selected as a research medium because it contains no phytosterols and consequently, no phytosterol oxides. GC results showed that no detectable level of -sitosterol oxides was present in the extracts. Moreover, the possibility of interconversion of each oxide during saponification and SPE purification was also evaluated. While each individual 20 g of -sitosterol oxide was separately spiked into 200 mg of butter and followed workup by the complete analytical method, no transformation happened between the oxides. Only the added oxide could be recovered in the GC analysis. Due to the analytes complexity in the food matrix, a cleanup step prior to GC analysis was mandatory. SiOH-SPE cartridges were used to isolate and concentrate polar lipids (containing phytosterol oxides) from low-polarity lipids and neutral lipid fraction (containing non-oxidized sterols) present in the unsaponifiable matter. In the work of Lampi et al. [20], an elution volume of 5 mL of hexane/diethyl ether (1:1, v/v) was used to remove the non-oxidized sterols from the cartridge prior to the oxide fraction. However, in the present study, this elution volume was reduced to 4 mL of cyclohexane/diethyl ether because trace amounts of 5,6␣epoxy and 5,6-epoxysitosterol were detected in the fifth
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milliliter of this elution. Ten replicate experiments and GC results showed no sterol oxides in the 4 mL of the cyclohexane/diethyl ether fraction (1:1, v/v), while all oxides were quantitatively removed from the SPE cartridge by the following 5 mL of acetone. 3.4. Formation of β-sitosterol oxides by heating β-sitosterol Phytosterols are basically stable compounds. However, under specific conditions, such as high temperature (>100 ◦ C) in the presence of air, oxidation process of phytosterols may occur in the same way as for cholesterol [2,24,38,39]. Some authors also reported the effect of refining and storage on the ppm level of the sterol oxides formed [31,33,40]. In order to study the effect of high temperatures on the formation of sterol oxides, 1 mg of pure -sitosterol was heated to 100, 150, or 200 ◦ C. No detectable level of oxides was found at 100 ◦ C in 30 min. At 150 ◦ C and 200 ◦ C, the main -sitosterol oxides represented during heating were 7␣-hydroxy, 7-hydroxy, 5,6␣-epoxy, 5,6-epoxy, and 7ketositosterol. No 5␣,6-dihydroxysitosterol was formed certainly due to the lack of water in the system. The major oxide formed at the two temperatures was 7-ketositosterol. The formation rate was quite constant during the first 30 min of heating at 150 ◦ C (Fig. 2(a)). In contrast, when -sitosterol was heated at 200 ◦ C, the oxide content increased very quickly during the first 15 min, then slightly decreased after 20–25 min. This phenomenon was probably due to thermodegradation and evaporation of the oxides formed (Fig. 2(b)). Similar results were shown in previous reports for the thermooxidation of stigmasterol or cholesterol [20,39]. 3.5. Quantification of β-sitosterol oxides in oils Vegetable oils or plant sterol-enriched foods are complex media because they contain several different phytosterols. -Sitosterol is among the most representative phytosterols, along with campesterol, stigmasterol, and other various types of minor phytosterols. The initial content of -sitosterol in the oils used was quantified by applying the analytical method to 200 mg of oil samples, and analyzing the fraction eluted with cyclohexane/diethyl ether (1:1, v/v) after spiking with 20 g of cholestanol as an internal standard. The total amounts of phytosterols were 2.6 and 1.6 mg/g in sunflower and olive oils, and the concentrations of -sitosterol were 1.5 and 0.9 mg/g in the oils, respectively. As mentioned above, the phytosterols contained in vegetable oils are labile through thermolytic and oxidation reactions when subjected to high temperatures or frying conditions. Fig. 3 presents the typical SIM chromatograms of the polar fractions in thermo-oxidized oils. The peaks of oxides were identified by their retention times as well as mass spectra, and quantitated with their characteristic ions of SIM mode along with their RRF, respectively, as shown in Tables 1 and 2.
Fig. 2. Major oxides formed by heating -sitosterol at different temperatures. Each -sitosterol oxide was quantified by capillary GC–MS with SIM mode: (a) 150 ◦ C, (b) 200 ◦ C. () 7␣-hydroxysitosterol; () 7hydroxysitosterol; () 5,6-epoxysitosterol; (×) 5,6␣-epoxysitosterol; () 7-ketositosterol.
Tables 4 and 5 summarize the concentration of six sitosterol oxides formed before and after heating of sunflower and olive oils at 150 and 200 ◦ C for 60 min. Initially, 7␣-hydroxy (6.5 g/g of oil), 7-hydroxy (7.0 g/g of oil), 7-keto (35.2 g/g of oil), and traces of 5␣,6dihydroxysitosterol (2.1 g/g of oil) were found in the native sunflower oil analyzed before heating. These oxides may be formed during oil processing and refinement [33,40]. No detectable levels of sterol oxides were found in the original olive oil. With heating, the amount of oxides formed in sunflower oil increased slowly during the first 30 min at 150 ◦ C and the first 15 min at 200 ◦ C. The total content was about 75 g/g of oil. After these durations, the content of total and individual -sitosterol oxides increased quickly at both temperatures with a total level about three times higher at 200 ◦ C (815.1 g/g of oil) than that at 150 ◦ C (240.9 g/g of oil) after 60 min. The content of 5␣,6-dihydroxysitosterol (2–10 g/g) did not change significantly at two temperatures. In olive oil, minor amounts of the oxides were formed after heating the olive oil at 150 ◦ C for 30 min or at 200 ◦ C for
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Fig. 3. Typical selection ion monitoring (SIM) chromatograms of m/z 356 + 353 + 485 + 413 + 395 + 432 + 396 obtained by GC–MS analysis of (a) six sitosterol oxides and internal standards (20 g of each compound), as well as in (b) the polar fractions of the heated sunflower oil, and in (c) the polar fraction of the heated olive oil. The upper chromatograms are corresponding total ion chromatogram (TIC) of oxides standards and polar fractions in heated oils. Chromatographic peaks: (1) cholestanol (internal standard); (2) 19-hydroxycholesterol (internal standard); (3) 7␣-hydroxysitosterol; (4) 7-hydroxysitosterol; (5) 5,6-epoxysitosterol; (6) 5,6␣-epoxysitosterol; (7) 5␣,6-dihydroxysitosterol; (8) 7-ketositosterol.
904
Table 4 Concentrations of -sitosterol oxides (g/g of oil) after heating sunflower oil at 150 or 200 ◦ C for different durationsa Analytes
150 ◦ C
200 ◦ C
0 min
5 min
15 min
30 min
60 min
0 min
5 min
15 min
30 min
60 min
7␣-Hydroxysitosterol 7-Hydroxysitosterol 5,6-Epoxysitosterol 5,6␣-Epoxysitosterol 5␣,6-Dihydroxysitosterol 7-Ketositosterol
6.5 ± 0.3 7.0 ± 0.3 n.d. n.d. 2.1 ± 0.4 35.2 ± 2.4
9.7 ± 0.6 9.6 ± 0.7 2.1 ± 0.2 1.2 ± 0.0 1.8 ± 0.1 41.7 ± 1.3
9.8 ± 0.2 9.4 ± 0.3 2.9 ± 0.2 2.0 ± 0.2 1.8 ± 0.0 44.0 ± 1.1
10.3 ± 0.4 10.2 ± 0.5 3.1 ± 0.6 1.1 ± 0.1 2.0 ± 0.2 45.9 ± 2.2
31.6 ± 2.3 49.5 ± 5.0 45.0 ± 4.3 24.3 ± 4.8 3.9 ± 0.6 86.6 ± 8.3
6.5 ± 0.3 7.0 ± 0.3 n.d. n.d. 2.1 ± 0.4 35.2 ± 2.4
10.1 ± 0.4 9.7 ± 0.5 2.9 ± 0.2 1.1 ± 0.0 1.8 ± 0.0 41.8 ± 1.0
10.6 ± 0.3 11.5 ± 0.4 2.2 ± 0.2 1.2 ± 0.1 2.1 ± 0.2 49.4 ± 0.7
33.1 ± 3.7 46.2 ± 3.8 46.9 ± 2.3 20.2 ± 2.2 3.8 ± 0.4 72.9 ± 1.4
81.3 ± 9.3 98.9 ± 8.7 220.3 ± 11.2 147.3 ± 21.5 9.1 ± 0.8 258.2 ± 16.5
Total amount (g/g oil) % -sitosterol oxidized
50.8 2.0
66.1 2.3
69.9 2.4
72.6 2.6
240.9 9.5
50.8 2.0
67.4 2.4
77.0 2.7
223.1 7.6
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n.d. = not detected. a Mean ± standard derivation (n = 3).
Table 5 Concentration of -sitosterol oxides (g/g oil) after heating olive oil at 150 or 200 ◦ C for different durationsa Analytes
150 ◦ C
200 ◦ C
0 min
5 min
15 min
30 min
60 min
0 min
5 min
15 min
30 min
60 min
7␣-Hydroxysitosterol 7-Hydroxysitosterol 5,6-Epoxysitosterol 5,6␣-Epoxysitosterol 5␣,6-Dihydroxysitosterol 7-Ketositosterol
n.d. n.d. n.d. n.d. n.d. n.d.
0.5 ± 0.0 0.5 ± 0.0 n.d. n.d. n.d. 0.7 ± 0.1
0.6 ± 0.1 0.6 ± 0.1 n.d. n.d. n.d. 1.0 ± 0.0
0.9 ± 0.1 1.1 ± 0.2 1.5 ± 0.3 1.0 ± 0.1 n.d. 1.5 ± 0.2
5.4 ± 0.2 9.5 ± 0.2 8.0 ± 0.4 3.8 ± 0.3 0.4 ± 0.3 10.0 ± 0.3
n.d. n.d. n.d. n.d. n.d. n.d.
0.5 ± 0.1 0.6 ± 0.2 n.d. n.d. n.d. 0.7 ± 0.2
0.8 ± 0.1 1.1 ± 0.2 0.6 ± 0.2 0.2 ± 0.3 n.d. 1.4 ± 0.2
21.1 ± 2.4 32.6 ± 3.0 31.5 ± 1.8 17.7 ± 0.2 2.3 ± 0.0 20.5 ± 1.0
50.3 ± 1.1 54.8 ± 1.2 94.5 ± 5.7 56.6 ± 5.6 3.2 ± 0.5 105.9 ± 9.3
Total amount (g/g oil) % -sitosterol oxidized
n.d. 0
1.7 1.0
2.2 1.3
6.0 3.0
37.1 7.2
n.d. 0
1.8 1.1
4.1 2.4
125.7 4.8
365.3 20.3
n.d. = not detected. a Mean ± standard derivation (n = 3).
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15 min, but they contributed to less than 3% of the -sitosterol oxidized. Conversely, the sunflower oil already contained similar levels of these oxides in its initial state. Trace levels of 5␣,6-dihydroxysitosterol were found in olive oil (<4 g/g) after heating. The result also showed that higher temperature will lead to more oxides formed. The total amounts of -sitosterol oxides in olive oil were 37.1 and 365.3 g/g of oil for heating at 150 and 200 ◦ C after 60 min, respectively. In the present heating assay of -sitosterol and oils, the  configuration of the oxides formed more favorably than that of the corresponding ␣ oxides. This result was in agreement with the previous report that formation of the -epimer of cholesterol epoxide was favored over the ␣-epimer [6]. The main oxides formed were the same at 150 and 200 ◦ C in the two oils. At 150 ◦ C, 7-hydroxy and 7-ketositosterol were present in the highest concentrations, while at 200 ◦ C, 7-ketositosterol quantitatively predominated, followed by 5,6-epoxysitosterol. As the effect of cooking time on the oxidation of cholesterol in meats [36], the formation rate of phytosterol oxides in plant oils differed according to the heating time and temperatures. Lower temperature and shorter periods might avoid the rapid formation of phytosterol oxides. However, significant amounts of oxides were detected in the oils heated at 200 ◦ C for 60 min (20–25% of -sitosterol oxidized). The total content of -sitosterol oxides formed at 200 ◦ C for 60 min with sunflower oil was about two times higher than that of olive oil under the same conditions.
Acknowledgements The authors thank Dr. Jennifer Wytko for her help in the preparation of the manuscript. This work was supported by a grant from the Minist`ere de la Jeunesse, de l’Education Nationale et de la Recherche, France (RARE 015 No. 02 P 0640).
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