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A highly sensitive quantification of phytosterols through an inexpensive derivatization
Chemistry and Physics of Lipids 166 (2013) 18–25
Contents lists available at SciVerse ScienceDirect
Chemistry and Physics of Lipids journal homepage: www.elsevier.com/locate/chemphyslip
A highly sensitive quantification of phytosterols through an inexpensive derivatization Songbai Liu ∗ , Huina Ruan Department of Food Science and Nutrition, Zhejiang University, 866 Yuhangtang Road, Hangzhou 310058, China
a r t i c l e
i n f o
Article history: Received 9 October 2012 Received in revised form 29 November 2012 Accepted 7 December 2012 Available online 19 December 2012 Keywords: Phytosterol Derivatization HPLC Quantification Detection Determination
a b s t r a c t A highly sensitive method for quantification of phytosterols based on HPLC has been developed by derivatization with the benzoyl chromophore. Introduction of the chromophore, benzoyl group, to phytosterols via simple and inexpensive derivatization greatly improved the UV response at 254 nm. Quantification of phytosterols was effectively performed by HPLC analysis with methyl benzoate as the internal standard after derivatization. This new method demonstrated outstanding yield of recovery (>95%) and excellent sensitivity (ng level) and was applicable for sterols from either plant or animal sources. This method is generally useful in phytosterol studies. © 2013 Elsevier Ireland Ltd. All rights reserved.
1. Introduction Phytosterols are plant-derived natural compounds that share structural similarity with cholesterol that is the predominant sterol in animals including humans. Increasing animal and human studies have revealed that high intakes of phytosterols can improve serum lipid profiles and reduce the risk of cardiovascular disease (Kendall and Jenkins, 2004) and caners such as breast and prostate cancer (Ju et al., 2004; Awad et al., 2001, 2000). Phytosterols are present in all plant foods, but the best dietary sources are unrefined plant oils including vegetable, nut, and olive oils (Ostlund, 2002) as well as nuts, seeds, whole grains, and legumes (De Jong et al., 2003). Due to their established beneficial effects, a vast number of research projects have been initiated to study the detailed molecular mechanism of biological activities of phytosterols and their derivatives especially phytosterol esters (St-Onge and Jones, 2003; Moruisi et al., 2006; Ellegard et al., 2007; Van Horn et al., 2008; Grundy, 2005). Scientists in food industry have shown growing interest in introduction of phytosterols in the modern diet (Berger et al., 2004; Volpe et al., 2001; Ostlund, 2004; Jauhiainen et al., 2006; Allen et al., 2008; Devaraj et al., 2004, 2006). During these studies, it is prerequisite to quantify phytosterols. Many methods have been developed for quantitative analysis
∗ Corresponding author. Tel.: +86 571 88982186; fax: +86 571 88982186. E-mail address: [email protected] (S. Liu). 0009-3084/$ – see front matter © 2013 Elsevier Ireland Ltd. All rights reserved. http://dx.doi.org/10.1016/j.chemphyslip.2012.12.002
since the discovery of phytosterols. The digitonin precipitation method emerged as the first generation approach and used to be frequently applied (IUPAC, 1979). Then a more sensitive colorimetric method was introduced which involved enzymatic oxidation of phytosterols and soon became obsolete owing to tedious operation and expensive reagents (Naudet and Hautfenne, 1985). At present the modern analytical methods, GC (Song et al., 2008; Kamm et al., 2002; Verleyen et al., 2001) and HPLC (Breinholder et al., 2002; Careri et al., 2001; Sanchez-Machado et al., 2004), have been employed in the determination of phytosterols. However, high boiling point of phytosterols necessitates heavy sample preparation and results in limited scope during GC analysis. In addition, expensive detectors including evaporative light scattering detector (ELSD) (Honda et al., 2010; Nair et al., 2006) and mass spectrometry (MS) (Canabate-Diaz et al., 2007) are usually required during HPLC analysis because of their low intensity of UV absorption. Therefore, developing a sensitive method with simple operation and inexpensive equipments for phytosterol quantification is highly desirable. Attempt to a highly sensitive method based on HPLC was initiated in our lab since HPLC generally has much broader scope of samples particularly for those of high boiling point. In this paper the progress of our study was disclosed. The high sensitivity was achieved by introduction of a chromophore to phytosterols through an instant derivatization of the hydroxyl group with inexpensive reagents. The derivatized phytosterols have strong UV absorption and can be selectively detected by the common UV detector without disturbance from impurities with short-wavelength UV absorption
S. Liu, H. Ruan / Chemistry and Physics of Lipids 166 (2013) 18–25
that is always a problem for non-derivatized phytosterols. To minimize errors in operation, an internal standard method was applied. Very good reproducibility and high yield of recovery have been achieved employing an inexpensive internal standard. Thus the newly developed method guarantees highly sensitive and selective detection of phytosterols with inexpensive reagents and simple operation and is generally useful in phytosterol studies. The details in development of this method are described as follows. 2. Materials and methods
19
2.5. HPLC analysis The HPLC system (Shimadzu, Japan) consisted of a high-pressure pump, an injector, a normal phase column (Hypersil SiO2 , 4.6 mm i.d. × 250 mm, 5 m, Elite), a column oven, and a UV detector. The mobile phase was hexane/isopropyl alcohol (99:1, v/v). The column was maintained at 35 ◦ C in the column oven. The flow rate was 1.0 mL/min. The detection wavelength was 254 nm. The injection volume was 20 L. Each sample was run three times and the average of the area of UV absorption was used for analysis.
2.1. Material and reagents
2.6. Calibration curve preparation
Phytosterol (30% stigmasterol, 5% brassicasterol, 20% campesterol and 40% -sitosterol, total purity 95%) were purchased from Jiatian Biotechnology (Xi’an, China). Sodium bicarbonate, anhydrous magnesium sulfate, and tetrahydrofuran were obtained from Sinopharm Chemical Reagent (China). Cholesterol, benzoyl chloride, methyl benzoate, pyridine, and ethylenediamine were obtained from Aladdin Reagent (China). Hexane and isopropyl alcohol (HPLC grade) were obtained from Tjshield Fine Chemicals (Hebei, China). All chemicals were of analytical grade unless otherwise noted.
Quantification of phytosterols was accomplished through a calibration curve prepared by HPLC through the following procedure. A stock solution of benzoyl phytosterol standard was prepared by dissolving 44.5 mg of benzoyl phytosterol standard into 10 mL hexane calibrated with a 10 mL volumetric flask. A stock solution of methyl benzoate was prepared by dissolving 116.6 mg methyl benzoate into 100 mL hexane calibrated with a 100 mL volumetric flask. Nine samples (9 calibration points) were prepared by mixing the stock solution of benzoyl phytosterol standard with that of methyl benzoate over the range of 5:1–1:5 (phytosterol ester:methyl benzoate, v/v) for HPLC analysis. The linear regression equation between the relative area of UV absorption and relative molarity for benzoyl phytosterol standard and the internal standard (methyl benzoate) was established by fitting the corresponding calibration points.
2.2. Phytosterol purification The commercially available phytosterols of 95% purity was further purified according to a modified literature procedure (Chen and Wu, 2004). To phytosterol (10 g, purity 95%) in a 500 mL Erlenmeyer flask was added 150 mL absolute ethanol. The flask capped with a stopper was heated at 80 ◦ C in a water bath water until all the phytosterols were dissolved in ethanol. Then heating was stopped. The flask was allowed to stand in the water bath and cooled down naturally to the ambient temperature. The precipitated phytosterols as a white solid were collected by filtration and washed 2 times by ethanol. The afforded phytosterols was recrystallized again with 50 mL absolute ethanol following the same procedure as above. The third recrystallization was performed instead with 150 mL hexane and heated at 60 ◦ C and provided the refined phytosterols for use. 2.3. Phytosterol derivatization Phytosterols were derivatized with benzoyl chloride following the procedure described below. To a solution of physterols in tetrahydrofuran was added 2 equivalent benzoyl choride and pyridine. The reaction mixture was stirred at ambient temperature. The progress of the reaction was monitored by TLC. The reaction was stopped as soon as phytosterols were completely consumed (typically 1 h). 2.4. Benzoyl phytosterol standard preparation Benzoyl phytosterol standard was prepared via the general derivatizing procedure after purification. Upon completion of the derivatization the resultant residue was dissolved in chloroform and transferred to a separatory funnel. The organic phase was washed 3 times by 1 N hydrochloric acid, followed by aqueous sodium bicarbonate. Then the organic phase was dried over anhydrous magnesium sulfate. The solvent was evaporated under reduced pressure. The afforded residue was further purified by flash chromatography (silica gel, petroleum ether/ethyl acetate = 20:1, v/v) to give benzoyl phytosterol standard for use in HPLC. The identity of benzoyl phytosterol standard was further confirmed by 1 H NMR spectroscopy.
2.7. Phytosterol quantification by HPLC analysis after derivatization Upon completion of the derivatization the reaction mixture was added ethylenediamine (two equivalent of the applied benzoyl chloride) and stirred further for around 2 min to destroy residual benzoyl chloride. The reaction mixture was diluted with hexane and transferred to a separatory funnel. The organic phase was washed 3 times by 1 N hydrochloric acid followed by aqueous sodium bicarbonate. Then the organic phase was dried over anhydrous magnesium sulfate. After filtration a specified quantity of the internal standard (methyl benzoate) was added in the filtrate and used for HPLC analysis. The quantification was performed on the basis of the established linear regression equation. Yield of recovery for the phytosterols was calculated according to the following equation: Yield of recovery (%) =
W2 W1 × Mp = × 100% W0 Mbp × w0
W0 , the original amount of phytosterols taken for the detection; W1 , the detected amount of benzoyl phytosterols by HPLC analysis according to the established linear regression equation; W2 , the detected amount of phytosterols calculated from W1 ; Mp, the molecular weight of phytosterols, 414.71 was used based on the major component of -sitosterol; Mbp, the molecular weight of benzoyl phytosterols, 518.83 was used based on the major component of -sitosterol. 3. Results and discussion 3.1. Phytosterol derivatization Phytosterol has a characteristic secondary hydroxyl group that can be readily esterified and etherified. To achieve strong UV absorption at 254 nm for phytosterols benzoyl group was chosen as the chromophore. Benzoyl chloride was employed to introduce the benzoyl group to phytosterols because of its remarkably high
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S. Liu, H. Ruan / Chemistry and Physics of Lipids 166 (2013) 18–25
Alkyl
Alkyl benzoyl chloride pyridine tetrahydrofuran
HO
O Ph
phytosterols
O benzoyl phytosterols
Fig. 1. Derivatization of phytosterols by benzoyl chloride.
(b)
Alkyl
O Ph
O
Signals from the benzoyl group
8.0
7.5
7.0
6.5
6.0
5.5
5.0 4.5 4.0 3.5 ChemicalShift (ppm)
3.0
2.5
2.0
1.5
1.0
0.5
2.5
2.0
1.5
1.0
0.5
(a)
Alkyl
HO
8.0
Fig. 2.
1
7.5
7.0
6.5
6.0
5.5
5.0 4.5 4.0 3.5 ChemicalShift(ppm)
3.0
H NMR spectra of (a) phytosterols and (b) benzoyl phytosterols. The introduction of benzoyl group was indicated by the signals in the spectrum (b).
S. Liu, H. Ruan / Chemistry and Physics of Lipids 166 (2013) 18–25
Fig. 3. HPLC analysis (UV detector, 254 nm) of (a) benzoyl phytosterols, (b) methyl benzoate and (c) a mixture of benzoyl phytosterols and methyl benzoate.
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S. Liu, H. Ruan / Chemistry and Physics of Lipids 166 (2013) 18–25
0.12
8
100
6 5 4 3
A1/A2 = 1.3552(n1/n2) - 0.0217
2
R2 = 0.997
1 0 0
1
2
3
4
5
6
95 0.08 90 0.04 85
The molar ratio of benzoyl phytosterols to methyl benzoate (n1/n2)
0
80 1
Fig. 4. Calibration curve of benzoyl phytosterols to methyl benzoate. The horizontal axis and the vertical axis are the molar ratio (n1 /n2 ) and relative intensity of UV absorption (A1 /A2 , 254 nm) of benzoyl phytosterols to methyl benzoate.
Yield of recovery (%)
7
Mass of phytosterols (g)
Relative intensity of UV absorption (A1/A2)
22
2
3
4
Detected mass
5
6
Loaded mass
7
8
9
10
Yield of recovery
Fig. 5. Yield of recovery for quantification of phytosterols over the range of 20–110 mg phytosterols.
3.2. Quantification of phytosterols via derivatization by HPLC-UV Internal standard method is the first option to eliminate any potential systematic errors instead of external standard method. The internal standard chosen for a specific compound to be quantified should fulfill a few requirements: easily separable with the compound, similar UV response to the compound, chemically stable. Thus methyl benzoate was chosen as the internal standard since it is stable and easily separable and has the same chromophore with benzoyl phytosterols. Intensive studies revealed the optimum condition for HPLC analysis. Under the optimum condition (hexane/isopropyl alcohol = 99/1; 254 nm) benzoyl phytosterols (retention time = 3.14 min) were completely separated with methyl benzoate (retention time = 3.66 min) as shown in Fig. 3. Another advantage was the short retention time (3–4 min) reducing the time cost. The calibration curve of relative UV response to relative molarity for benzoyl phytosterols and methyl benzoate was obtained at different relative molarity levels over the range of 0.2–5.0 (9 calibration points). Least squares regression of the calibration points produced the linear regression equation (Fig. 4) with good linearity (regression coefficient, r = 0.997) that laid a solid ground for quantification of phytosterols. Finally the absolute quantity of phytosterols was calculated from the quantity of benzoyl phytosterols derived from the linear regression equation. 3.3. Recovery efficacy for the quantification To test the reliability of this method, recovery of variable quantity of phytosterols over the range of 20–110 mg was investigated (Fig. 5). When the quantity of phytosterols for detection was less than 60 mg variable yields of recovery were observed. The gap between quantity of recovery and authentic quantity resulted from
the loss during the extraction step in the derivatization of phytosterols. However, high yield of recovery for a small quantity of sample still can be achieved once sufficient extraction operation is performed as exemplified by the result of 30 mg phytosterols (yield of recovery ≈ 95%). When larger quantities (≥60 mg) of phytosterols were applied, high yields of recovery (>95%) were generally realized. Hence this method is reliable and accurate for quantification of phytosterols even with the amount of tens of milligrams. To further test the efficacy of this newly developed method, cholesterol, the predominant sterol in animals, was examined. The result exhibited excellent yield of recovery (99.7%) when about 30 mg of cholesterol was used for detection (Fig. 6), which revealed generality of this method. It was suggested that this method was generally useful for detection of sterols from either plant or animal sources.
3.4. Detection limit of quantification After establishment of the reliability and reproducibility of the new method, study on the detection limit was conducted (Fig. 7). Initially a solution of benzoyl phytosterols with the concentration of 4.45 g/L was prepared. Loading 20 L of the solution that corresponded to 89 g of benzoyl phytosterols gave a strong UV response of 951 mV. Then the solution was diluted by 10 times. The UV response for the loading of 8.9 g of benzoyl phytosterols was 75 mV. Further dilution by 10 times demonstrated a UV response of 12 mV for 0.89 g sample. Finally 89 ng of benzoyl phytosterols still produced a good UV response of 2 mV with excellent signal-tonoise ratio. These results exhibited this method was very sensitive and the detection limit reached to ng level.
0.04
Mass of cholesterol (g)
reactivity with the hydroxyl group which ensured thorough derivatization of phytosterols. Phytosterols treated with excess amount of benzoyl chloride and pyridine in tetrahydrofuran realized full derivatization in a couple of hours at ambient temperature (Fig. 1). Upon completion of the derivatization the reaction mixture was easily worked up for further HPLC analysis. Excess benzoyl chloride was instantly scavenged by ethylenediamine and the resultant byproducts and pyridine were cleanly washed out by aqueous hydrochloric acid. The afforded benzoyl phytosterols were further isolated and characterized. As shown in Fig. 2, 1 H NMR of benzoyl phytosterols indicated the characteristic benzoyl group compared with that of phytosterols. Therefore, strong UV absorption, inexpensive reagents (benzoyl chloride and pyridine) and simple operation contribute to the usefulness of the newly developed derivatization method.
0.0317 g
0.0318 g
0.03
0.02
0.01
0 Detected mass
Loaded mass
Fig. 6. Recovery for quantification of cholesterol (yield of recovery = 99.7%).
S. Liu, H. Ruan / Chemistry and Physics of Lipids 166 (2013) 18–25
Fig. 7. HPLC response of different quantities of benzoyl phytosterols: (a) 89 g, (b) 8.9 g, (c) 0.89 g and (d) 89 ng.
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S. Liu, H. Ruan / Chemistry and Physics of Lipids 166 (2013) 18–25
Fig. 7. Continued)
4. Conclusion A highly sensitive method for quantification of phytosterols was developed. The pending issue of low UV absorption for phytosterols was resolved. Introduction of the chromophore, benzoyl group, to phytosterols via simple and inexpensive derivatization greatly improved the UV response at 254 nm. Quantification of phytosterols was effectively performed by HPLC analysis with methyl benzoate as the internal standard after derivatization. This new method demonstrated outstanding yield of recovery (>95%) and excellent sensitivity (ng level) and was applicable for sterols from either plant or animal sources. This method is generally useful in phytosterol studies. Acknowledgements This work was supported by Zijin Plan of Zhejiang University and ‘The Fundamental Research Funds for the Central Universities’. References Allen, R.R., Carson, L., Kwik-Uribe, C., Evans, E.M., Erdman Jr., J.W., 2008. Daily consumption of a dark chocolate containing flavanols and added sterol esters affects cardiovascular risk factors in a normotensive population with elevated cholesterol. Journal of Nutrition 138, 725–731. Awad, A.B., Downie, A., Fink, C.S., Kim, U., 2000. Dietary phytosterol inhibits the growth and metastasis of MDA-MB-231 human breast cancer cells grown in SCID mice. Anticancer Research 20, 821–824. Awad, A.B., Fink, C.S., Williams, H., Kim, U., 2001. In vitro and in vivo (SCID mice) effects of phytosterols on the growth and dissemination of human prostate cancer PC-3 cells. European Journal of Cancer Prevention 10, 507–513. Berger, A., Jones, P.J., Abumweis, S.S., 2004. Plant sterols: factors affecting their efficacy and safety as functional food ingredients. Lipids Health Disorder 3, 5–23. Breinholder, P., Mosca, L., Lindner, W., 2002. Concept of sequential analysis of free and conjugated phytosterols in different plant matrices. Journal of Chromatography 777, 67–82. Canabate-Diaz, B., Carretero, A.S., Fernandez-Gutierrez, A., Vega, A.B., Frenich, A.G., Vidal, J.L.M., et al., 2007. Separation and determination of sterols in olive oil by HPLC-MS. Food Chemistry 102, 593–598. Careri, M., Elviri, L., Mangia, A., 2001. Liquid chromatography-UV determination and liquid chromatography-atmospheric pressure chemical ionization mass spectrometric characterization of sitosterol and stigmasterol in soybean oil. Journal of Chromatography A 935, 249–257. Chen, M., Wu, M., 2004. Preparation of mixed plant sterol standard sample and its application in analysis. Food Science 25, 125–128. De Jong, A., Plat, J., Mensink, R.P., 2003. Metabolic effects of plant sterols and stanols. Journal of Nutritional Biochemistry 14, 362–369.
Devaraj, S., Jialal, I., Vega-Lopez, S., 2004. Plant sterol-fortified orange juice effectively lowers cholesterol levels in mildly hypercholesterolemic healthy individuals. Arteriosclerosis, Thrombosis, and Vascular Biology 24, 25–28. Devaraj, S., Autret, B.C., Jialal, I., 2006. Reduced-calorie orange juice beverage with plant sterols lowers C-reactive protein concentrations and improves the lipid profile in human volunteers. American Journal of Clinical Nutrition 84, 756–761. Ellegard, L.H., Andersson, S.W., Normen, A.L., Andersson, H.A., 2007. Dietary plant sterols and cholesterol metabolism. Nutrition Reviews 65, 39–45. Grundy, S.M., 2005. Stanol esters as a component of maximal dietary therapy in the national cholesterol education program adult treatment panel III report. American Journal of Cardiology 96, 47–50. Honda, A., Miyazaki, T., Ikegami, T., Iwamoto, J., Yamashita, K., et al., 2010. Numazawa M, Highly sensitive and specific analysis of sterol profiles in biological samples by HPLC–ESI–MS/MS. Journal of Steroid Biochemistry and Molecular Biology 121, 556–564. IUPAC, 1979. Standard Methods for the Analysis of Oils, Fats and Derivatives, sixth ed., Pergamon Press, Oxford. Jauhiainen, T., Salo, P., Niittynen, L., Poussa, T., Korpela, R., 2006. Effects of low-fat hard cheese enriched with plant stanol esters on serum lipids and apolipoprotein B in mildly hypercholesterolaemic subjects. European Journal of Clinical Nutrition 60, 1253–1257. Ju, Y.H., Clausen, L.M., Allred, K.F., 2004. Almada AL and Helferich WG, Beta-sitosterol, beta-sitosterol glucoside, and a mixture of beta-sitosterol and beta-sitosterol glucoside modulate the growth of estrogen-responsive breast cancer cells in vitro and in ovariectomized athymic mice. Journal of Nutrition 134, 1145–1151. Kamm, W., Dionisi, F., Hischenhuber, C., Schmarr, H.G., Engel, K.H., 2002. Rapid detection of vegetable oils in milk fat by on-line LC-GC analysis of betasitosterol as marker. European Journal of Lipid Science and Technology 104, 756–761. Kendall, C.W., Jenkins, D.J., 2004. A dietary portfolio: maximal reduction of lowdensity lipoprotein cholesterol with diet. Current Atherosclerosis Reports 6, 492–498. Moruisi, K.G., Oosthuizen, W., Opperman, A.M., 2006. Phytosterols/stanols lower cholesterol concentrations in familial hypercholesterolemic subjects: a systematic review with meta-analysis. Journal of the American College of Nutrition 25, 41–48. Nair, V.D.P., Kanfer, I., Hoogmartens, J., 2006. Determination of stigmasterol, sitosterol and stigmastanol in oral dosage forms using high performance liquid chromatography with evaporative light scattering detection. Journal of Pharmaceutical and Biomedical Analysis 41, 731–737. Naudet, M., Hautfenne, A., 1985. Determination of total sterols in fats and oils. Pure and Applied Chemistry 57, 899–904. Ostlund Jr., R.E., 2002. Phytosterols in human nutrition. Annual Review of Nutrition 22, 533–549. Ostlund Jr., R.E., 2004. Phytosterols and cholesterol metabolism. Current Opinion in Lipidology 15, 37–41. Sanchez-Machado, D.I., Lopez-Hernandez, J., Paseiro-Losada, P., Lopez-Cervantes, J., 2004. An HPLC method for the quantification of sterols in edible seaweeds. Biomedical Chromatography 18, 183–190. Song, Z., Peng, Y., Liu, F., 2008. Gas chromatography analytical of mixed phytosterols. Guangzhou Chemical Industry 36, 72–74.
S. Liu, H. Ruan / Chemistry and Physics of Lipids 166 (2013) 18–25 St-Onge, M.P., Jones, P.J., 2003. Phytosterols and human lipid metabolism: efficacy, safety, and novel foods. Lipids 38, 367–375. Van Horn, L., McCoin, M., Kris-Etherton, P.M., Burke, F., Carson, J.A.S., Champagne, C.M., et al., 2008. The evidence for dietary prevention and treatment of cardiovascular disease. Journal of the American Dietetic Association 108, 287–331.
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Volpe, R., Niittynen, L., Korpela, R., Sirtori, C., Bucci, A., Fraone, N., et al., 2001. Effects of yoghurt enriched with plant sterols on serum lipids in patients with moderate hypercholesterolaemia. British Journal of Nutrition 86, 233–239. Verleyen, T., Verhe, R., Garcia, L., Dewettinck, K., Huyghebaert, A., De Greyt, W., 2001. Gas chromatographic characterization of vegetable oil deodorization distillate. Journal of chromatographic 921, 277–285.
Contents lists available at SciVerse ScienceDirect
Chemistry and Physics of Lipids journal homepage: www.elsevier.com/locate/chemphyslip
A highly sensitive quantification of phytosterols through an inexpensive derivatization Songbai Liu ∗ , Huina Ruan Department of Food Science and Nutrition, Zhejiang University, 866 Yuhangtang Road, Hangzhou 310058, China
a r t i c l e
i n f o
Article history: Received 9 October 2012 Received in revised form 29 November 2012 Accepted 7 December 2012 Available online 19 December 2012 Keywords: Phytosterol Derivatization HPLC Quantification Detection Determination
a b s t r a c t A highly sensitive method for quantification of phytosterols based on HPLC has been developed by derivatization with the benzoyl chromophore. Introduction of the chromophore, benzoyl group, to phytosterols via simple and inexpensive derivatization greatly improved the UV response at 254 nm. Quantification of phytosterols was effectively performed by HPLC analysis with methyl benzoate as the internal standard after derivatization. This new method demonstrated outstanding yield of recovery (>95%) and excellent sensitivity (ng level) and was applicable for sterols from either plant or animal sources. This method is generally useful in phytosterol studies. © 2013 Elsevier Ireland Ltd. All rights reserved.
1. Introduction Phytosterols are plant-derived natural compounds that share structural similarity with cholesterol that is the predominant sterol in animals including humans. Increasing animal and human studies have revealed that high intakes of phytosterols can improve serum lipid profiles and reduce the risk of cardiovascular disease (Kendall and Jenkins, 2004) and caners such as breast and prostate cancer (Ju et al., 2004; Awad et al., 2001, 2000). Phytosterols are present in all plant foods, but the best dietary sources are unrefined plant oils including vegetable, nut, and olive oils (Ostlund, 2002) as well as nuts, seeds, whole grains, and legumes (De Jong et al., 2003). Due to their established beneficial effects, a vast number of research projects have been initiated to study the detailed molecular mechanism of biological activities of phytosterols and their derivatives especially phytosterol esters (St-Onge and Jones, 2003; Moruisi et al., 2006; Ellegard et al., 2007; Van Horn et al., 2008; Grundy, 2005). Scientists in food industry have shown growing interest in introduction of phytosterols in the modern diet (Berger et al., 2004; Volpe et al., 2001; Ostlund, 2004; Jauhiainen et al., 2006; Allen et al., 2008; Devaraj et al., 2004, 2006). During these studies, it is prerequisite to quantify phytosterols. Many methods have been developed for quantitative analysis
∗ Corresponding author. Tel.: +86 571 88982186; fax: +86 571 88982186. E-mail address: [email protected] (S. Liu). 0009-3084/$ – see front matter © 2013 Elsevier Ireland Ltd. All rights reserved. http://dx.doi.org/10.1016/j.chemphyslip.2012.12.002
since the discovery of phytosterols. The digitonin precipitation method emerged as the first generation approach and used to be frequently applied (IUPAC, 1979). Then a more sensitive colorimetric method was introduced which involved enzymatic oxidation of phytosterols and soon became obsolete owing to tedious operation and expensive reagents (Naudet and Hautfenne, 1985). At present the modern analytical methods, GC (Song et al., 2008; Kamm et al., 2002; Verleyen et al., 2001) and HPLC (Breinholder et al., 2002; Careri et al., 2001; Sanchez-Machado et al., 2004), have been employed in the determination of phytosterols. However, high boiling point of phytosterols necessitates heavy sample preparation and results in limited scope during GC analysis. In addition, expensive detectors including evaporative light scattering detector (ELSD) (Honda et al., 2010; Nair et al., 2006) and mass spectrometry (MS) (Canabate-Diaz et al., 2007) are usually required during HPLC analysis because of their low intensity of UV absorption. Therefore, developing a sensitive method with simple operation and inexpensive equipments for phytosterol quantification is highly desirable. Attempt to a highly sensitive method based on HPLC was initiated in our lab since HPLC generally has much broader scope of samples particularly for those of high boiling point. In this paper the progress of our study was disclosed. The high sensitivity was achieved by introduction of a chromophore to phytosterols through an instant derivatization of the hydroxyl group with inexpensive reagents. The derivatized phytosterols have strong UV absorption and can be selectively detected by the common UV detector without disturbance from impurities with short-wavelength UV absorption
S. Liu, H. Ruan / Chemistry and Physics of Lipids 166 (2013) 18–25
that is always a problem for non-derivatized phytosterols. To minimize errors in operation, an internal standard method was applied. Very good reproducibility and high yield of recovery have been achieved employing an inexpensive internal standard. Thus the newly developed method guarantees highly sensitive and selective detection of phytosterols with inexpensive reagents and simple operation and is generally useful in phytosterol studies. The details in development of this method are described as follows. 2. Materials and methods
19
2.5. HPLC analysis The HPLC system (Shimadzu, Japan) consisted of a high-pressure pump, an injector, a normal phase column (Hypersil SiO2 , 4.6 mm i.d. × 250 mm, 5 m, Elite), a column oven, and a UV detector. The mobile phase was hexane/isopropyl alcohol (99:1, v/v). The column was maintained at 35 ◦ C in the column oven. The flow rate was 1.0 mL/min. The detection wavelength was 254 nm. The injection volume was 20 L. Each sample was run three times and the average of the area of UV absorption was used for analysis.
2.1. Material and reagents
2.6. Calibration curve preparation
Phytosterol (30% stigmasterol, 5% brassicasterol, 20% campesterol and 40% -sitosterol, total purity 95%) were purchased from Jiatian Biotechnology (Xi’an, China). Sodium bicarbonate, anhydrous magnesium sulfate, and tetrahydrofuran were obtained from Sinopharm Chemical Reagent (China). Cholesterol, benzoyl chloride, methyl benzoate, pyridine, and ethylenediamine were obtained from Aladdin Reagent (China). Hexane and isopropyl alcohol (HPLC grade) were obtained from Tjshield Fine Chemicals (Hebei, China). All chemicals were of analytical grade unless otherwise noted.
Quantification of phytosterols was accomplished through a calibration curve prepared by HPLC through the following procedure. A stock solution of benzoyl phytosterol standard was prepared by dissolving 44.5 mg of benzoyl phytosterol standard into 10 mL hexane calibrated with a 10 mL volumetric flask. A stock solution of methyl benzoate was prepared by dissolving 116.6 mg methyl benzoate into 100 mL hexane calibrated with a 100 mL volumetric flask. Nine samples (9 calibration points) were prepared by mixing the stock solution of benzoyl phytosterol standard with that of methyl benzoate over the range of 5:1–1:5 (phytosterol ester:methyl benzoate, v/v) for HPLC analysis. The linear regression equation between the relative area of UV absorption and relative molarity for benzoyl phytosterol standard and the internal standard (methyl benzoate) was established by fitting the corresponding calibration points.
2.2. Phytosterol purification The commercially available phytosterols of 95% purity was further purified according to a modified literature procedure (Chen and Wu, 2004). To phytosterol (10 g, purity 95%) in a 500 mL Erlenmeyer flask was added 150 mL absolute ethanol. The flask capped with a stopper was heated at 80 ◦ C in a water bath water until all the phytosterols were dissolved in ethanol. Then heating was stopped. The flask was allowed to stand in the water bath and cooled down naturally to the ambient temperature. The precipitated phytosterols as a white solid were collected by filtration and washed 2 times by ethanol. The afforded phytosterols was recrystallized again with 50 mL absolute ethanol following the same procedure as above. The third recrystallization was performed instead with 150 mL hexane and heated at 60 ◦ C and provided the refined phytosterols for use. 2.3. Phytosterol derivatization Phytosterols were derivatized with benzoyl chloride following the procedure described below. To a solution of physterols in tetrahydrofuran was added 2 equivalent benzoyl choride and pyridine. The reaction mixture was stirred at ambient temperature. The progress of the reaction was monitored by TLC. The reaction was stopped as soon as phytosterols were completely consumed (typically 1 h). 2.4. Benzoyl phytosterol standard preparation Benzoyl phytosterol standard was prepared via the general derivatizing procedure after purification. Upon completion of the derivatization the resultant residue was dissolved in chloroform and transferred to a separatory funnel. The organic phase was washed 3 times by 1 N hydrochloric acid, followed by aqueous sodium bicarbonate. Then the organic phase was dried over anhydrous magnesium sulfate. The solvent was evaporated under reduced pressure. The afforded residue was further purified by flash chromatography (silica gel, petroleum ether/ethyl acetate = 20:1, v/v) to give benzoyl phytosterol standard for use in HPLC. The identity of benzoyl phytosterol standard was further confirmed by 1 H NMR spectroscopy.
2.7. Phytosterol quantification by HPLC analysis after derivatization Upon completion of the derivatization the reaction mixture was added ethylenediamine (two equivalent of the applied benzoyl chloride) and stirred further for around 2 min to destroy residual benzoyl chloride. The reaction mixture was diluted with hexane and transferred to a separatory funnel. The organic phase was washed 3 times by 1 N hydrochloric acid followed by aqueous sodium bicarbonate. Then the organic phase was dried over anhydrous magnesium sulfate. After filtration a specified quantity of the internal standard (methyl benzoate) was added in the filtrate and used for HPLC analysis. The quantification was performed on the basis of the established linear regression equation. Yield of recovery for the phytosterols was calculated according to the following equation: Yield of recovery (%) =
W2 W1 × Mp = × 100% W0 Mbp × w0
W0 , the original amount of phytosterols taken for the detection; W1 , the detected amount of benzoyl phytosterols by HPLC analysis according to the established linear regression equation; W2 , the detected amount of phytosterols calculated from W1 ; Mp, the molecular weight of phytosterols, 414.71 was used based on the major component of -sitosterol; Mbp, the molecular weight of benzoyl phytosterols, 518.83 was used based on the major component of -sitosterol. 3. Results and discussion 3.1. Phytosterol derivatization Phytosterol has a characteristic secondary hydroxyl group that can be readily esterified and etherified. To achieve strong UV absorption at 254 nm for phytosterols benzoyl group was chosen as the chromophore. Benzoyl chloride was employed to introduce the benzoyl group to phytosterols because of its remarkably high
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S. Liu, H. Ruan / Chemistry and Physics of Lipids 166 (2013) 18–25
Alkyl
Alkyl benzoyl chloride pyridine tetrahydrofuran
HO
O Ph
phytosterols
O benzoyl phytosterols
Fig. 1. Derivatization of phytosterols by benzoyl chloride.
(b)
Alkyl
O Ph
O
Signals from the benzoyl group
8.0
7.5
7.0
6.5
6.0
5.5
5.0 4.5 4.0 3.5 ChemicalShift (ppm)
3.0
2.5
2.0
1.5
1.0
0.5
2.5
2.0
1.5
1.0
0.5
(a)
Alkyl
HO
8.0
Fig. 2.
1
7.5
7.0
6.5
6.0
5.5
5.0 4.5 4.0 3.5 ChemicalShift(ppm)
3.0
H NMR spectra of (a) phytosterols and (b) benzoyl phytosterols. The introduction of benzoyl group was indicated by the signals in the spectrum (b).
S. Liu, H. Ruan / Chemistry and Physics of Lipids 166 (2013) 18–25
Fig. 3. HPLC analysis (UV detector, 254 nm) of (a) benzoyl phytosterols, (b) methyl benzoate and (c) a mixture of benzoyl phytosterols and methyl benzoate.
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0.12
8
100
6 5 4 3
A1/A2 = 1.3552(n1/n2) - 0.0217
2
R2 = 0.997
1 0 0
1
2
3
4
5
6
95 0.08 90 0.04 85
The molar ratio of benzoyl phytosterols to methyl benzoate (n1/n2)
0
80 1
Fig. 4. Calibration curve of benzoyl phytosterols to methyl benzoate. The horizontal axis and the vertical axis are the molar ratio (n1 /n2 ) and relative intensity of UV absorption (A1 /A2 , 254 nm) of benzoyl phytosterols to methyl benzoate.
Yield of recovery (%)
7
Mass of phytosterols (g)
Relative intensity of UV absorption (A1/A2)
22
2
3
4
Detected mass
5
6
Loaded mass
7
8
9
10
Yield of recovery
Fig. 5. Yield of recovery for quantification of phytosterols over the range of 20–110 mg phytosterols.
3.2. Quantification of phytosterols via derivatization by HPLC-UV Internal standard method is the first option to eliminate any potential systematic errors instead of external standard method. The internal standard chosen for a specific compound to be quantified should fulfill a few requirements: easily separable with the compound, similar UV response to the compound, chemically stable. Thus methyl benzoate was chosen as the internal standard since it is stable and easily separable and has the same chromophore with benzoyl phytosterols. Intensive studies revealed the optimum condition for HPLC analysis. Under the optimum condition (hexane/isopropyl alcohol = 99/1; 254 nm) benzoyl phytosterols (retention time = 3.14 min) were completely separated with methyl benzoate (retention time = 3.66 min) as shown in Fig. 3. Another advantage was the short retention time (3–4 min) reducing the time cost. The calibration curve of relative UV response to relative molarity for benzoyl phytosterols and methyl benzoate was obtained at different relative molarity levels over the range of 0.2–5.0 (9 calibration points). Least squares regression of the calibration points produced the linear regression equation (Fig. 4) with good linearity (regression coefficient, r = 0.997) that laid a solid ground for quantification of phytosterols. Finally the absolute quantity of phytosterols was calculated from the quantity of benzoyl phytosterols derived from the linear regression equation. 3.3. Recovery efficacy for the quantification To test the reliability of this method, recovery of variable quantity of phytosterols over the range of 20–110 mg was investigated (Fig. 5). When the quantity of phytosterols for detection was less than 60 mg variable yields of recovery were observed. The gap between quantity of recovery and authentic quantity resulted from
the loss during the extraction step in the derivatization of phytosterols. However, high yield of recovery for a small quantity of sample still can be achieved once sufficient extraction operation is performed as exemplified by the result of 30 mg phytosterols (yield of recovery ≈ 95%). When larger quantities (≥60 mg) of phytosterols were applied, high yields of recovery (>95%) were generally realized. Hence this method is reliable and accurate for quantification of phytosterols even with the amount of tens of milligrams. To further test the efficacy of this newly developed method, cholesterol, the predominant sterol in animals, was examined. The result exhibited excellent yield of recovery (99.7%) when about 30 mg of cholesterol was used for detection (Fig. 6), which revealed generality of this method. It was suggested that this method was generally useful for detection of sterols from either plant or animal sources.
3.4. Detection limit of quantification After establishment of the reliability and reproducibility of the new method, study on the detection limit was conducted (Fig. 7). Initially a solution of benzoyl phytosterols with the concentration of 4.45 g/L was prepared. Loading 20 L of the solution that corresponded to 89 g of benzoyl phytosterols gave a strong UV response of 951 mV. Then the solution was diluted by 10 times. The UV response for the loading of 8.9 g of benzoyl phytosterols was 75 mV. Further dilution by 10 times demonstrated a UV response of 12 mV for 0.89 g sample. Finally 89 ng of benzoyl phytosterols still produced a good UV response of 2 mV with excellent signal-tonoise ratio. These results exhibited this method was very sensitive and the detection limit reached to ng level.
0.04
Mass of cholesterol (g)
reactivity with the hydroxyl group which ensured thorough derivatization of phytosterols. Phytosterols treated with excess amount of benzoyl chloride and pyridine in tetrahydrofuran realized full derivatization in a couple of hours at ambient temperature (Fig. 1). Upon completion of the derivatization the reaction mixture was easily worked up for further HPLC analysis. Excess benzoyl chloride was instantly scavenged by ethylenediamine and the resultant byproducts and pyridine were cleanly washed out by aqueous hydrochloric acid. The afforded benzoyl phytosterols were further isolated and characterized. As shown in Fig. 2, 1 H NMR of benzoyl phytosterols indicated the characteristic benzoyl group compared with that of phytosterols. Therefore, strong UV absorption, inexpensive reagents (benzoyl chloride and pyridine) and simple operation contribute to the usefulness of the newly developed derivatization method.
0.0317 g
0.0318 g
0.03
0.02
0.01
0 Detected mass
Loaded mass
Fig. 6. Recovery for quantification of cholesterol (yield of recovery = 99.7%).
S. Liu, H. Ruan / Chemistry and Physics of Lipids 166 (2013) 18–25
Fig. 7. HPLC response of different quantities of benzoyl phytosterols: (a) 89 g, (b) 8.9 g, (c) 0.89 g and (d) 89 ng.
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Fig. 7. Continued)
4. Conclusion A highly sensitive method for quantification of phytosterols was developed. The pending issue of low UV absorption for phytosterols was resolved. Introduction of the chromophore, benzoyl group, to phytosterols via simple and inexpensive derivatization greatly improved the UV response at 254 nm. Quantification of phytosterols was effectively performed by HPLC analysis with methyl benzoate as the internal standard after derivatization. This new method demonstrated outstanding yield of recovery (>95%) and excellent sensitivity (ng level) and was applicable for sterols from either plant or animal sources. This method is generally useful in phytosterol studies. Acknowledgements This work was supported by Zijin Plan of Zhejiang University and ‘The Fundamental Research Funds for the Central Universities’. References Allen, R.R., Carson, L., Kwik-Uribe, C., Evans, E.M., Erdman Jr., J.W., 2008. Daily consumption of a dark chocolate containing flavanols and added sterol esters affects cardiovascular risk factors in a normotensive population with elevated cholesterol. Journal of Nutrition 138, 725–731. Awad, A.B., Downie, A., Fink, C.S., Kim, U., 2000. Dietary phytosterol inhibits the growth and metastasis of MDA-MB-231 human breast cancer cells grown in SCID mice. Anticancer Research 20, 821–824. Awad, A.B., Fink, C.S., Williams, H., Kim, U., 2001. In vitro and in vivo (SCID mice) effects of phytosterols on the growth and dissemination of human prostate cancer PC-3 cells. European Journal of Cancer Prevention 10, 507–513. Berger, A., Jones, P.J., Abumweis, S.S., 2004. Plant sterols: factors affecting their efficacy and safety as functional food ingredients. Lipids Health Disorder 3, 5–23. Breinholder, P., Mosca, L., Lindner, W., 2002. Concept of sequential analysis of free and conjugated phytosterols in different plant matrices. Journal of Chromatography 777, 67–82. Canabate-Diaz, B., Carretero, A.S., Fernandez-Gutierrez, A., Vega, A.B., Frenich, A.G., Vidal, J.L.M., et al., 2007. Separation and determination of sterols in olive oil by HPLC-MS. Food Chemistry 102, 593–598. Careri, M., Elviri, L., Mangia, A., 2001. Liquid chromatography-UV determination and liquid chromatography-atmospheric pressure chemical ionization mass spectrometric characterization of sitosterol and stigmasterol in soybean oil. Journal of Chromatography A 935, 249–257. Chen, M., Wu, M., 2004. Preparation of mixed plant sterol standard sample and its application in analysis. Food Science 25, 125–128. De Jong, A., Plat, J., Mensink, R.P., 2003. Metabolic effects of plant sterols and stanols. Journal of Nutritional Biochemistry 14, 362–369.
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