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Enzymatic synthesis of phytosteryl lipoate and its antioxidant properties
Food Chemistry 240 (2018) 736–742
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Enzymatic synthesis of phytosteryl lipoate and its antioxidant properties ⁎
MARK
Huiqi Wang, Chengsheng Jia , Xue Xia, Eric Karangwa, Xiaoming Zhang State Key Laboratory of Food Science and Technology, School of Food Science and Technology, Jiangnan University, Wuxi 214122, Jiangsu, PR China
A R T I C L E I N F O
A B S T R A C T
Chemical compounds studied in this article include: Phytosterols (PubChem CID: 12303662) Lipoic acid (PubChem CID: 6112) Stigmasterol (PubChem CID: 5280794)
In this work, an enzymatic route for synthesizing phytosteryl lipoate was successfully set up for the first time. The structure of final product phytosteryl lipoate was determined by Fourier Transform Infrared (FTIR), Mass Spectrometry (MS) and Nuclear Magnetic Resonance (NMR). The maximum conversion of 71.2% was achieved when the following conditions were employed: 150 mmol/L phytosterol, 1: 2.5 M ratio of phytosterol to lipoic acid, 10 g/L of 4 Å molecular sieves and 60 g/L Candida rugosa in 2-methyl-2-butanol/n-hexane (1:1, v/v) at 55 °C for 96 h. The physicochemical properties including solubility and antioxidant ability of phytosteryl lipoate in oil were assessed. The results revealed that phytosteryl lipoate possessed over twice as much oil solubility as free phytosterol and also showed better antioxidant ability. Investigation on its biological functions will be the main object in the future work.
Keywords: Phytosteryl lipoate Enzymatic synthesis Oil solubility Peroxide value Antioxidant activity
1. Introduction Phytosterol plays important roles in pharmaceutical industries because of the vast physiological significances such as cholesterol-lowering, antioxidant and anticancer functions (Alappat, Valerio, & Awad, 2010; Bard, Paillard, & Lecerf, 2015; Danesi, Gomez-Caravaca, de Biase, Verardo, & Bordoni, 2016; Woyengo, Ramprasath, & Jones, 2009). In addition, plant sterols are one of healthy factors in food (Baumgartner, Mensink, & Plat, 2016; Dong et al., 2016; te Velde et al., 2015). However, the practical application of phytosterol is limited due to its poor solubility in both water and oil, which is caused by its special chemical structure. Consequently, there is a need to modify phytosterol structure with its functions retained. Up to now, most studies have focused on the esterification of phytosterol with fatty acids to improve its solubility, however, there are a few reports on phytosteryl esters with their improved biological functions (Fu et al., 2014; Tan & Shahidi, 2012, 2013). What is noteworthy is that these phytosteryl esters can be decomposed into two initial reactants after digestion (Carden, Hang, Dussault, & Carr, 2015; Lubinus, Barnsteiner, Skurk, Hauner, & Engel, 2013). As a result, it’s crucial to choose a suitable reactant which can improve respectively the physicochemical properties and can also perform their beneficial functions together. Antioxidants have been always attracting the attention of many researchers, since they can prevent not only the oxidation of fat rich food to prolong their storage time but also senescence and some other diseases caused by reactive oxygen species (ROS) and free radicals in
⁎
Corresponding author. E-mail address: [email protected] (C. Jia).
http://dx.doi.org/10.1016/j.foodchem.2017.08.025 Received 31 May 2017; Received in revised form 30 July 2017; Accepted 4 August 2017 Available online 04 August 2017 0308-8146/ © 2017 Published by Elsevier Ltd.
body, such as Alzheimer’s disease (Rosini et al., 2005). Lipoic acid is a very important kind of vitamin B compounds and mainly presents in the liver tissue of animals and some plants such as tomato, broccoli, carrot and spinach. The high electron density in closed five-membered ring imparts lipoic acid significant electrophilic property and the ability to react with free radicals. In the body, lipoic acid can be reduced into dihydrogen lipoic acid which has two sulfydryls. Both lipoic acid and dihydrogen lipoic acid have good resistance to oxidation and can work together in fat soluble and water soluble environment. In this relation, they are widely known as natural universal antioxidants. Lipoic acid owns various useful functions for human health, which has been already used in medical treatment, such as the prevention of diabetes, cancer, cardiovascular disease and inflammation (Benchekroun et al., 2016; Derosa, D'Angelo, Romano, & Maffioli, 2016; Moura, de Andrade, dos Santos, & Goulart, 2015; Tsuji-Naito, Ishikura, Akagawa, & Saeki, 2010; Ying et al., 2010). Meanwhile, it is also extensively applied in cosmetics and health food fields (Leysen & Aerts, 2016; Merry, Kirk, & Goyns, 2008). Researchers have already reported the synthesis of lipoic acid conjugates for health applications (Hsieh et al., 2015; Kaki, Balakrishna, & Prasad, 2014; Lahiani, Hidmi, Katzhendler, Yavin, & Lazarovici, 2016). Recent study has demonstrated that the phytosterol/lipoic acid combination could work together and played a preferable and superior cholesterol-lowering role than phytosterol or lipoic acid alone (Rideout et al., 2016). Phytosteryl lipoate as shown in Fig. 1, synthesized through esterification of phytosterol with lipoic acid, was supposed to play better physiological functions with smaller
Food Chemistry 240 (2018) 736–742
H. Wang et al. O
Fig. 1. Synthetic route of phytosteryl lipoate. OH
R
S S
Lipoic acid
O
Lipase
+
O S S
R
R=
,
,
,
etc
Phytosteryl lipoate
HO R=
,
,
,
etc
Phytosterol
2.3. Purification of esterified reaction mixture
amounts than either phytosterol or lipoic acid. Enzymatic method is a great route for mild reaction, high safety and less by-products, which meets the requirements for good health and green. In recent years, lipases have been widely used for catalyzing esterification reactions of phytosterols. Schär and Nyström produced steryl ferulates using enzymatic method for the first time (Schär & Nyström, 2016). Miao et al. have selected Candida rugosa as catalyst to develop phytosteryl laurate in non-aqueous media (Miao et al., 2014). To the best of our knowledge, the enzymatic synthesis of phytosteryl lipoate has not still been well documented. The present work was therefore aimed to explore a lipase-catalyzed method to synthesize phytosteryl lipoate and investigate its optimal reaction conditions. After purification by Thin-Layer Chromatography (TLC) and silica gel column chromatography, final product’s chemical structure was determined by FT-IR, NMR and MS. Meanwhile, the solubility and antioxidant activity of phytosteryl lipoate were also evaluated.
After the esterification reaction, the mixture was filtered to remove the lipase and molecular sieves, and then the extract was treated with 0.1 M NaHCO3 aqueous solution to neutralize excess lipoic acid. The filtrates were evaporated by rotary evaporator to remove the solvents, then, 1.0 g of the dried sample was dissolved in petroleum ether (60–90 °C)/ethyl acetate/formic acid mixture (15:1:0.02, v/v/v). The obtained solutions were subsequently applied to a silica gel column (24 × 1200 mm) and eluted with petroleum ether (60–90 °C)/ethyl acetate/formic acid (15:1:0.02, v/v/v) at a flow rate of 0.5 mL/min. The eluate was detected by HPLC and collected at the same time, the esters were then obtained after the solvents evaporation by rotary evaporation. 2.4. HPLC analysis The samples (10 µL) taken out from reaction systems were injected in Waters 1525 HPLC analysis equipped with a symmetry-C18 column (5 μm, 4.6 × 150 mm, Waters, USA) at 45 °C. The sample was eluted with methanol (mobile phase) at a flow rate of 1 mL/min. The effluent was monitored with a Grace Alltech 3300 evaporative light scattering detector (ELSD) at 55 °C and nitrogen was used as the carrier gas at a flow rate of 1.5 L/min. The esterification rate of phytosterol with lipoic acid to form phytosteryl lipoate was calculated via calibration curve and purified phytosteryl lipoate was used as external standard. The conversion was defined as follows:
2. Materials and methods 2.1. Chemicals Phytosterols were a generous gift from Jiangsu Spring Fruit Biological Products Co., Ltd. (Taixing, P.R. China). The purity of plant sterols was > 97% (63% β-sitosterol and 37% stigmasterol). Lipoic acid was purchased from Xi’an Jinheng Chemical Co., Ltd. (Shanxi, China). Novozym 435 (lipase B from Candida antarctica, immobilized on a macroporous acrylic resin, 10,000 PLU/g), Lipozyme TL IM (lipase from Thermomyces lanuginosus, immobilized on silica granulation, 250 IUN/g), Lipozyme RM IM (lipase from Rhizomucor miehei, immobilized on an anionic exchange resin, 275 IUN/g) were obtained from Novo Nordisk Co., Ltd. (Shanghai, China). Candida rugosa (lyophilized powder, Type VII, 847 U/mg) was supplied by Sigma-Aldrich Co., Ltd. (Shanghai, China). Methanol of spectral grade was obtained from Tedia Company Inc. (Shanghai, China). The rapeseed oil was purchased from the local supermarket. The 3 Å and 4 Å molecular sieves and other common reagents of analytical grade were from Sinopharm Chemical Reagent Co., Ltd. (Shanghai, China).
XPSE (%) = CPSE /CPS × 100
(1)
where CPSE is the actual concentration of phytosteryl lipoate at the end of the reaction, CPS is the initial concentration of phytosterol. 2.5. FTIR analysis The purified phytosteryl lipoate was dried completely under vacuum and then mixed with KBr. FTIR analysis was conducted on a FT-IR spectrophotometer (Thermo Nicolet iS10 FTIR, Thermo Electron, USA) using an attenuated total reflectance method with the spectral scanning scope for 400–4000 cm−1, the number of scans is 32 and resolution is 4 cm−1.
2.2. Enzymatic synthesis of phytosteryl lipoate
2.6. MS analysis
Typically, phytosterol (1 mmol) and lipoic acid (1–4 mmol) were dissolved in 10 mL organic solvent at first, followed by the addition of 0–0.4 g of molecular sieves and lipase (0.4–0.6 g). The reaction was carried in a shaking water bath at 150 r/min, for 48–120 h at 40–60 °C. All samples were performed in triplicate.
Liquid chromatography mass spectrometry (Maldi Synapt Q-Tof, Waters, USA) was used to further identify the stigmasteryl ester under positive-ion electron spray ionization (ESI) mode. The MS parameters were listed as follows: capillary voltage, 3.5 kV; cone voltage, 30 V; source block temperature, 100 °C; desolvation temperature, 400 °C; 737
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3. Results and discussion 3.1. Structural analysis of phytosteryl lipoate 3.1.1. HPLC analysis The conversion and purity of the final product were determined by HPLC. At the end of the reaction, the reaction mixture was quantitatively analyzed by HPLC and the results were presented in Fig. S1. The peaks of phytosterol and lipoic acid were confirmed by their respective standards. The phytosterol used in the present experiment mainly contained stigmasterol and sitosterol, consequently, the synthetic phytosteryl esters were stigmasteryl lipoate and sitosteryl lipoate. According to the spectrogram and polarity analysis, lipoic acid was firstly eluted at the retention time of 1.745 min, and phytosterols (stigmasterol and sitosterol) were then eluted at the retention time of 3.128 and 3.374 min, respectively. Additionally, two new contiguous peaks at 6.064 min and 6.632 min appeared and these compounds might be stigmasteryl lipoate and sitosteryl lipoate, respectively. Fig. 2. FTIR spectra of lipoic acid (a) and phytosteryl lipoate (b).
3.1.2. FTIR analysis The IR spectra of lipoic acid and the newly synthesized product were compared, as shown in Fig. 2(a), the broad peak between 2400 cm−1 and 3300 cm−1 indicated the presence of eOH group from eCOOH. The strong peak at 1692.84 cm−1 was the signal of the C]O group from eCOOH. In Fig. 2(b), the peaks at 2898.38 cm−1 and 1366.80 cm−1 were the stretching vibration and bending vibrations of CeH groups for methylene groups, respectively. The peaks at 2866.94 cm−1 and 1462.24 cm−1 were the stretching vibration and bending vibrations of CeH groups for methyl groups. The strong peak at 1732.46 cm−1 was the signal of the C]O group, and the medium peaks at 1173.43 cm−1 and 1131.68 cm−1 were the stretching vibration of CeO group of ester bond. In summary, the disappeared absorption signal of free carboxyl group and the emerging ester bond provided evidence that the new product was phytosteryl lipoate.
desolvation gas flow, 700 lit/hr; cone gas flow rate, 50 l/h and scan range, m/z 50–1000. 2.7. NMR analysis NMR experiments were performed on an NMR spectrometer (Avance III 400 MHz, Bruker, Switzerland) using CDCl3 as the solvent and tetramethylsilane as the internal standard. The 1H and 13C NMR spectrum were acquired at 400.14 and 100.62 MHz at 298.0 K, respectively. 2.8. Determination of oil solubility Due to the complex oil components, the solubility of phytosterol and phytosteryl lipoate was determined by direct observation, and phytosteryl ferulate was also measured as a reference. The method was described as follows: 200 mg samples were put into a flask while stirred and heated to 30 °C, then rapeseed oil was added drop by drop until the all samples were entirely dissolved and the oil amount of consumption was noted down immediately. The oil solubility was represented as grams (g) per litre (L) at 30 °C.
3.1.3. MS analysis HPLC-MS with ESI was used to further confirm stigmasteryl lipoate under positive-ion mode. The main advantage of this method is that the reaction mixture could be used for analysis without separation and purification prior to analysis (He et al., 2014). Theoretically, the relative molecular weight for stigmasteryl lipoate was 600. As shown in Fig. 3, the molecular ion [M+H]+ at m/z 601.4 and a sodium adduct molecular ion [M+Na]+ at m/z 623.4 were also identified. The molecular ions [M+H]+ and [M+Na]+ were also observed in previous reports about phytosteryl esters using ESI-MS with positive-ion mode (He et al., 2012, 2016). In addition, a major fragment [M+H-lipoic acid]+ at m/z 395.4 was observed. Similar [M+H-ferulic acid]+ and [M+H-lauric acid]+ fragments have been also reported in the MS analysis of campesteryl ferulate and ergosterol laurate (He et al., 2014; Tan & Shahidi, 2011). These results proved that the synthesized product was stigmasteryl lipoate.
2.9. Antioxidant activity studies in oil The antioxidant ability of phytosteryl lipoate (purity, > 98%) in edible oil was measured by peroxide value (POV) method. It is wellknown that edible oils are full of various unsaturated fatty acids which are subjected to oxidation easily (Köckritz & Martin, 2008). The edible oils containing samples firstly dissolved in chloroform and glacial acetic acid, then the peroxide in the oils would generate iodine after reacting with potassium iodide, finally the amount of iodine was measured using the standard solution of sodium thiosulfate by titration and POV can be indirectly calculated. The specific method was described by Min et al. (2014) with slight modification according to GB/T5009.227-2016, and the POV was calculated using the following equation:
X1 = (V−V0) × C× 0.1269 × 100/m
3.1.4. NMR analysis The synthesized and purified stigmasteryl lipoate was analyzed by 1 H and 13C NMR. The results were listed as follows: 1H NMR (400 MHz, CDCl3): δ = 0.63 (3 H, s, 18-H), 0.73 (6 H, d, J = 7.2 Hz, 26–27-H), 0.78 (3 H, d, J = 6.4 Hz, 21-H), 0.89–0.98 (6 H, m), 0.95 (3 H, s, 19-H), 1.01–1.23 (6 H, m), 1.32–1.66 (16 H, m, 3′-H, 4′-H, 5′-H, etc.), 1.77–2.08 (5 H, m, 7′-Ha, etc.), 2.21–2.25 (4 H, m, 2′-H, etc.), 2.36–2.43 (1 H, dt, J = 4.0, 8.0 Hz, 7′-Hb), 3.02–3.15 (2 H, m, 8′-H), 3.47–3.54 (1 H, m, 6′-H), 4.52–4.58 (1 H, m, 3-H), 4.94 (1 H, dd, J = 8.0, 12.0 Hz, 22-H or 23-H), 5.08 (1 H, dd, J = 8.0, 12.0 Hz, 22-H or 23-H), 5.30 (1 H, d, J = 4.0, 6-H). 13C NMR (100 MHz, CDCl3): δ = 12.06 (29CH3), 12.26 (18-CH3), 18.99 (21-CH3), 19.34 (19-CH3), 21.03 (26CH3), 21.10 (27-CH3), 21.23 (CH2), 24.37 (11-CH2), 24.78 (3′-CH2),
(2)
The X1 represented POV (g/100 g), V was the volume (mL) of standard solution of sodium thiosulfate consumed by samples, while the volume of the sample blank was expressed as V0 (mL). C represented the concentration of sodium thiosulfate (0.002 mol/L), and m (g) corresponded to the mass of the samples. The number 0.1269 was the mass of iodine which was equal to 1.00 mL standard solution of sodium thiosulfate (Na2S2O3 = 1.00 mol/L) and 100 was the conversion coefficient. 738
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Fig. 3. HPLC-MS chromatogram of stigmasteryl lipoate.
25.42 (15-CH2), 27.83 (CH2), 28.74 (4′-CH2), 28.92 (CH2), 31.87 (8- or 25-CH), 31.89 (8- or 25-CH), 34.45 (5′-CH2), 34.61 (8′-CH2), 36.63 (10C), 37.01 (CH2), 38.17 (CH2), 38.49 (CH2), 39.64 (CH2), 40.23 (20-CH), 40.50 (7′-CH), 42.22 (13-C), 50.06 (9-CH), 51.24 (24-CH), 55.95 (17CH), 56.37 (6′-CH), 56.79 (14-CH), 73.87 (3-C), 122.63 (6-C), 129.34 (22-C), 138.31 (23-C), 139.72 (5-C), 172.91 (C = O). The 1H NMR and 13 C NMR spectra of phytosteryl lipoate were displayed in Figs. S2 and S3, respectively. The proton signals were overlapped at δ 0.63–3.15 in 1H NMR, the chemical shift below δ 1.2 could primarily be attributed to methyl groups in stigmasterol and those between δ 1.2 and 3.2 were ascribed to methylene in both stigmasterol and lipoic acid. The proton peaks of lipoic acid in the product were at δ 1.32–1.66, 1.77–2.08, 2.21–2.25, 2.36–2.43, 3.02–3.15 and 3.47–3.54, respectively. These results are greatly consistent with the findings reported by Madawala, Andersson, Jastrebova, Almeida, and Dutta (2012). The chemical shift of 3-position proton was at δ 3.5 in free phytosterol (He et al., 2016), while it shifted to δ 4.5 in the product due to the formation of ester bond, which agreed with its resonances in other phytosteryl esters (Kobayashi et al., 2014). Similarly, the 13C NMR analysis of the compound showed that the chemical shift of 3-C (δ 73.87) was lower than that in free stigmasterol (δ 71.8) (Huang, Cao, Xu, & Chen, 2011).
Table 1 Influence of different solvents and lipases on conversion. Solvent
tert-Butanol 2-Methyl-2-butanol n-Hexane Acetone/n-hexane (1:1, v/v) 2-Methyl-2-butanol/nhexane (1:1, v/v) tert-Butanol/n-hexane (1:1, v/v)
Conversion (%) Novozyme 435
Candida Rugosa
TL IM
RM IM
0.8 0.6 0.2 3.2 1.3
0.6 1.7 0.4 5.3 6.5
0.4 0.6 0.2 1.7 3.4
0.2 0.3 0.1 0.2 0.1
0.8
1.2
2.2
0.6
Reaction condition: 50 mmol/L phytosterol, 1.5:1 M ratio of lipoic acid to phytosterol, 20 g/L lipase and 40 g/L of 4 Å molecular sieve at 150 r/min and 45 °C for 72 h.
shown in Table 1. This could be attributed to the poor solubility of lipoic acid in n-hexane. On the contrary, though both substrates have good solubility in tert-butanol and 2-methyl-2-butanol, the deprivation of water made the enzyme inactive, hence the conversion was low. In such situation, mixed solvents are often used when the significant difference exists between the polarities of two substrates (Gu et al., 2014; Zhao et al., 2011). Thus, n-hexane was mixed with the three solvents of different polarities (1:1, v/v), respectively, and the results showed that the conversion was significantly improved in co-solvent system, especially in acetone/n-hexane and 2-methyl-2-butanol/n-hexane system, respectively. Four lipases in different forms were used in the present study. Novozyme 435, Lipozyme TL IM and Lipozyme RM IM are all immobilized lipases, while Candida rugosa are powder lipase. Different lipases, originated from diverse sources of microorganism, presented dramatically different catalytic efficiency for the esterification of phytosterol with lipoic acid. The catalytic efficiency of Lipozyme RM IM was the lowest for the reaction conversion. Novozyme 435 in acetone/ n-hexane (1:1, v/v) showed the highest conversion of 3.2% and
3.2. Lipase-catalyzed synthesis of phytosteryl lipoate 3.2.1. Screen of enzyme and solvent Different solvents, with log P values from −0.26 to 3.50, and several kinds of lipase are screened and their conversions are recorded in Table 1. The influences of solvent are mainly embodied in two aspects: on the one hand, the solubility of substrates is a pivotal factor for esterification rate, on the other hand, every enzyme needs a certain amount of water to maintain its configuration and activity, and the higher the polarity of solvent is, the lower the activity of enzyme is. nHexane is a commonly used solvent with low polarity and food safety, however, its conversion was the lowest among the selected solvents as 739
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Fig. 4. Influence of phytosterol concentration (a), molar ratio (b), molecular sieve (c), lipase load (d), temperature (e) and time (f) on esterification rate. (Reaction condition: (a) 1.5:1 M ratio of lipoic acid to phytosterol, 20 g/L lipase and 40 g/L of 4 Å molecular sieve, 45 °C, 72 h; (b) 150 mmol/L phytosterol, 20 g/L lipase and 40 g/L of 4 Å molecular sieve, 45 °C, 72 h; (c) 150 mmol/L phytosterol, 2.5:1 M ratio of lipoic acid to phytosterol and 20 g/L lipase, 45 °C, 72 h; (d) 150 mmol/L phytosterol, 2.5:1 M ratio of lipoic acid to phytosterol and 10 g/L of 4 Å molecular sieve, 45 °C, 72 h; (e) 150 mmol/L phytosterol, 2.5:1 M ratio of lipoic acid to phytosterol, 60 g/L lipase and 10 g/L of 4 Å molecular sieve, 72 h; (f) 150 mmol/L phytosterol, 2.5:1 M ratio of lipoic acid to phytosterol, 60 g/L lipase and 10 g/L of 4 Å molecular sieve, 55 °C.)
substrates and catalysis temperature used in this study. Kim and Akoh (2007) reported that Candida rugosa was a substrate-dependent enzyme for the synthesis of phytosterol esters. Hence, Candida rugosa in 2-methyl-2-butanol/n-hexane (1:1, v/v) was considered as the most desirable biocatalyst and solvent.
Lipozyme TL IM in 2-methyl-2-butanol/n-hexane (1:1, v/v) showed the maximum conversion of 3.4%. However, Candida rugosa in acetone/nhexane (1:1, v/v) and 2-methyl-2-butanol/n-hexane (1:1, v/v) showed the highest conversion of 5.3% and 6.5%, respectively. It was clear that Candida rugosa showed high catalytic ability compared to the other three immobilized enzymes at 45 °C. Though Novozyme 435 was regarded as the most active lipase in most cases, it could not achieve better conversion than C. Rugosa, which might be ascribed to the
3.2.2. Effect of substrate concentration The influence of different 740
phytosterol
concentrations
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3.2.7. Effect of reaction time As shown in Fig. 4(f), the esterification rate linearly increased with increasing reaction time from 24 h to 96 h, and the maximum conversion (71.2%) was reached after 96 h. When the reaction prolonged up to 120 h, the esterification rate decreased to 68.9%. Therefore, the reaction had reached equilibrium after around 96 h. According to the above optimization results, the maximum yield (> 71%) could be reached under the selected conditions: 150 mmol/L phytosterol, 1:2.5 M ratio of phytosterol to lipoic acid, 60 g/L Candida rugosa, 10 g/L 4Å molecular sieves in 2-methyl-2-butanol/n-hexane (1:1, v/v) at 55 °C for 96 h at a stirring rate of 150 r/min.
(25–200 mmol/L) on the esterification of phytosterol with lipoic acid was investigated and the results are shown in Fig. 4(a). At first, the conversion linearly increased with increasing substrate concentration, and the conversion reached the highest rate (10.2%) at 150 mol/L phytosterol concentration, then decreased afterward with further increasing concentration. The conversion dropped to 7.8%, this observation might be attributed to the restricted total solubility of substrates and reduced enzyme activity, because enzyme active sites were occupied by redundant substrates (He et al., 2010). 3.2.3. Effect of molar ratio Different molar ratios varied from 1:1.5 to 3:1 of lipoic acid to phytosterol were used in a series experiments. As presented in Fig. 4(b), it has been demonstrated that the excess of either phytosterol or lipoic acid was beneficial for esterification reaction, the conversion was lower (5.6%) at equimolar substrates. Our findings are in good agreement with previous studies (He et al., 2016; No, Zhao, Lee, Lee, & Kim, 2013). Considering the purification procedure, lipoic acid is more convenient to separate than phytosterol, therefore excess lipoic acid was used. As the molar ratio increased from 1:1 to 2.5:1, the esterification rate was boosted from 5.6% to 16.1%, then dropped to 11.3% when the molar ratio was 3:1. This might be attributed to the denaturation of enzyme in acidic condition.
3.3. Comparison of solubility In general, substances can become more hydrophobic with the increase of lipophilic group in the molecules, and their solubility in oils will accordingly be enhanced. The solubility of phytosteryl lipoate and phytosteryl ferulate reached 20.4 g/L and 22.5 g/L in rapeseed oil at 30 °C, respectively, while the solubility of phytosterols was only 9.1 g/L under the same condition. Conclusively, the esterification of phytosterol with lipoic acid improved its solubility in edible oils. 3.4. Antioxidant ability
3.2.4. Effect of molecular sieve concentration Controlling the water content in the reaction system is of great concern. Too much water could make reaction equilibrium move to the left side, while lack of water would have impact on the conformation and the catalytic efficiency of enzyme, then conversion would decline under any of the two situations. Hence, it is very critical to choose appropriate variety and quantity of dehydrating agent. Due to the economic factors and separation procedure, different dosage of 3 Å and 4 Å molecular sieves were used in our study. 3 Å molecular sieve has better ability to remove water compared with 4 Å molecular sieve (He et al., 2010). From Fig. 4(c), it could be concluded that molecular sieve possessed a large effect on esterification rate and the conversion using 4 Å molecular sieve was higher than that using 3 Å molecular sieve. The conversion reached the maximum value (30.2%) when the 4 Å molecular sieve concentration was 10 g/L and drastically declined afterward, while the conversion kept decreasing when 3 Å molecular sieve was used. The water in reaction system was gradually reduced with the increase concentration of molecular sieve, which led to the decline of enzyme activity. Therefore, 10 g/L of 4 Å molecular sieve was selected for further experiments.
The antioxidant ability of phytosteryl lipoate was evaluated by POV determination and was compared to that of phytosterol. Phytosteryl ferulate, the main ingredient of γ-oryzanol which has already been used commercially for its cholesterol-lowering, antioxidant and antiinflammation properties, was selected for reference. All samples were analyzed in triplicate. The same amount (0.2 g/kg) of different samples were added into rapeseed oil and their corresponding POVs were shown in Fig. 5. Since oil samples were continuously heated at 60 °C, all samples would generate more and more peroxide and their POVs increased. According to GB2716-2005, POV should be below 0.25 g/ 100 g in oil. As denoted in Fig. 5, the rapeseed oil without any additives showed the highest POVs, while the POVs of oil samples containing phytosteryl lipoate showed the lowest POVs from 24 h to 84 h. The POVs of blank sample were 0.23 g/100 g at 60 h and 0.27 g/100 g at 72 h. The samples treated with phytosterol had slightly lower POVs, and its POVs were 0.21 g/100 g at 60 h and 0.25 g/100 g at 72 h. It could be seen that both phytosteryl lipoate and phytosteryl ferulate had better oxidation resistance than phytosterol, their POVs were 0.19 g/ 100 g and 0.22 g/100 g at 72 h, 0.23 g/100 g and 0.24 g/100 g at 84 h, respectively. The POVs of phytosteryl lipoate was lower than that of phytosteryl ferulate as a whole, which indicates that phytosteryl lipoate could potentially be used as an antioxidant in oils based food.
3.2.5. Effect of lipase load Fig. 4(d) illustrated the effect of different concentration of Candida rugosa varying from 20 g/L to 80 g/L on the conversion. As displayed in Fig. 4(d), the conversion increased from 28.9% to 38.4% with increasing Candida rugosa load from 20 g/L to 40 g/L, then reached to 47.2% when the lipase loading was 60 g/L. As the loading continued to increase up to 80 g/L, the esterification rate was slightly changed. Out of the economic aspects, 60 g/L lipase loading was selected for the further studies. 3.2.6. Effect of reaction temperature The substrates solubility and enzyme activity are both strongly dependent on the reaction temperature. The effect of reaction temperature (40 °C to 60 °C) on esterification rate was studied. As presented in Fig. 4(e), the initial esterification rate was 41.8% at 40 °C, and the conversion gradually increased with increasing temperature and reached the apex (56.2%) at 55 °C, then decreased with further increasing of the reaction temperature from 55 °C to 60 °C. These findings are in accordance with the results reported by Kim and Akoh (2007), which stipulated that Candida rugosa could play its best catalytic role when the reaction temperature was set at 55 °C.
Fig. 5. Peroxide value (POV) (g/100 g) of rapeseed oil treated with phytosterol, phytosteryl lipoate and phytosteryl ferulate at 60 °C.
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Enzymatic synthesis and characterization of 1-lipoyl-2-palmitoyl phosphatidylcholine: A novel phospholipid containing lipoic acid. European Journal of Lipid Science and Technology, 116, 1347–1353. Kim, B. H., & Akoh, C. C. (2007). Modeling and optimization of lipase-catalyzed synthesis of phytosteryl esters of oleic acid by response surface methodology. Food Chemistry, 102, 336–342. Kobayashi, T., Ogino, A., Miyake, Y., Mori, H., Hosoda, A., Fujita, M., ... Adachi, S. (2014). Lipase-catalyzed esterification of triterpene alcohols and phytosterols with oleic acid. Journal of the American Oil Chemists' Society, 91, 1885–1890. Köckritz, A., & Martin, A. (2008). Oxidation of unsaturated fatty acid derivatives and vegetable oils. European Journal of Lipid Science and Technology, 110, 812–824. Lahiani, A., Hidmi, A., Katzhendler, J., Yavin, E., & Lazarovici, P. (2016). Novel synthetic PEGylated conjugate of alpha-lipoic acid and tempol reduces cell death in a neuronal PC12 clonal line subjected to ischemia. ACS Chemical Neuroscience, 7, 1452–1462. Leysen, J., & Aerts, O. (2016). Further evidence of thioctic acid (alpha-lipoic acid) being a strong cosmetic sensitizer. Contact Dermatitis, 74, 182–184. Lubinus, T., Barnsteiner, A., Skurk, T., Hauner, H., & Engel, K. (2013). Fate of dietary phytosteryl/-stanyl esters: analysis of individual intact esters in human feces. European Journal of Nutrition, 52, 997–1013. Madawala, S., Andersson, R., Jastrebova, J., Almeida, M., & Dutta, P. (2012). Phytosterol and α-lipoic acid conjugates: synthesis, free radical scavenging capacity and RP-LCMS-APCI analysis. Polish Journal of Food and Nutrition Sciences, 62, 159–169. Merry, B. J., Kirk, A., & Goyns, M. H. (2008). Dietary lipoic acid supplementation can mimic or block the effect of dietary restriction on life span. Mechanisms of Ageing and Development, 129, 341–348. Miao, M., Liu, H., Jiang, B., Yang, C., Xia, X., & Zhang, T. (2014). Enzyme-catalysed synthesis of plant steryl laurate in non-aqueous media using salt hydrate pairs and its characterisation. Journal of Functional Foods, 7, 452–461. Min, Z., Guo, Z., Wang, K., Zhang, A., Li, H., & Fang, Y. (2014). Antioxidant effects of grape vine cane extracts from different Chinese grape varieties on edible oils. Molecules, 19, 15213–15223. Moura, F. A., de Andrade, K. Q., dos Santos, J. C. F., & Goulart, M. O. F. (2015). Lipoic acid: its antioxidant and anti-inflammatory role and clinical applications. Current Topics in Medicinal Chemistry, 15, 458–483. No, D. S., Zhao, T., Lee, J., Lee, J. S., & Kim, I. H. (2013). Synthesis of phytosteryl ester containing pinolenic acid in a solvent-free system using immobilized Candida rugosa lipase. Journal of Agricultural and Food Chemistry, 61, 8934–8940. Rideout, T. C., Carrier, B., Wen, S., Raslawsky, A., Browne, R. W., & Harding, S. V. (2016). Complementary cholesterol-lowering response of a phytosterol/alpha-lipoic acid combination in obese zucker rats. Journal of Dietary Supplements, 13, 283–299. Rosini, M., Andrisano, V., Bartolini, M., Bolognesi, M. L., Hrelia, P., Minarini, A., ... Melchiorre, C. (2005). Rational approach to discover multipotent anti-Alzheimer drugs. Journal of Medicinal Chemistry, 48, 360–363. Schär, A., & Nyström, L. (2016). Enzymatic synthesis of steryl ferulates. European Journal of Lipid Science and Technology, 118, 1557–1565. Tan, Z., & Shahidi, F. (2011). Chemoenzymatic synthesis of phytosteryl ferulates and evaluation of their antioxidant activity. Journal of Agricultural and Food Chemistry, 59, 12375–12383. Tan, Z., & Shahidi, F. (2012). A novel chemoenzymatic synthesis of phytosteryl caffeates and assessment of their antioxidant activity. Food Chemistry, 133, 1427–1434. Tan, Z., & Shahidi, F. (2013). Phytosteryl sinapates and vanillates: chemoenzymatic synthesis and antioxidant capacity assessment. Food Chemistry, 138, 1438–1447. te Velde, A. A., Brull, F., Heinsbroek, S. E., Meijer, S. L., Lutjohann, D., Vreugdenhil, A., & Plat, J. (2015). Effects of dietary plant sterols and stanol esters with low- and high-fat diets in chronic and acute models for experimental colitis. Nutrients, 7, 8518–8531. Tsuji-Naito, K., Ishikura, S., Akagawa, M., & Saeki, H. (2010). Alpha-Lipoic acid induces collagen biosynthesis involving prolyl hydroxylase expression via activation of TGFbeta-Smad signaling in human dermal fibroblasts. Connective Tissue Research, 51, 378–387. Woyengo, T. A., Ramprasath, V. R., & Jones, P. J. (2009). Anticancer effects of phytosterols. European Journal of Clinical Nutrition, 63, 813–820. Ying, Z. K., Kherada, N., Farrar, B., Kampfrath, T., Chung, Y. C., Simonetti, O., ... Sun, Q. H. (2010). Lipoic acid effects on established atherosclerosis. Life Sciences, 86, 95–102. Zhao, H., Zhang, Y., Lu, F., Bie, X., Lu, Z., & Ning, H. (2011). Optimized enzymatic synthesis of ascorbyl esters from lard using Novozym 435 in co-solvent mixtures. Journal of Molecular Catalysis B: Enzymatic, 69, 107–111.
4. Conclusion In this study, the phytosteryl lipoate was successfully synthesized by a direct enzymatic approach. The effects of 8 parameters (solvent, lipase, phytosterol concentration, molar ratio, molecular sieve, lipase load, temperature and time) on conversion were evaluated and the highest conversion was 71.2%. After conjugation with lipoic acid, the oil solubility and antioxidant ability of phytosterol have been greatly improved. This novel ester broadens the application of phytosterol in food and pharmaceutical industry and may also possess physiological functions of phytosterol and lipoic acid, such as cholesterol-lowering ability. Acknowledgement This research was financially supported by the Natural Science Foundation of Jiangsu Province (BK20161133) and program of “Collaborative Innovation Center of Food Safety and Quality Control in Jiangsu Province”. . Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.foodchem.2017.08.025. References Alappat, L., Valerio, M., & Awad, A. B. (2010). Effect of vitamin D and beta-sitosterol on immune function of macrophages. International Immunopharmacology, 10, 1390–1396. Bard, J. M., Paillard, F., & Lecerf, J. M. (2015). Effect of phytosterols/stanols on LDL concentration and other surrogate markers of cardiovascular risk. Diabetes Metabolism, 41, 69–75. Baumgartner, S., Mensink, R. P., & Plat, J. (2016). Effects of a plant sterol or stanol enriched mixed meal on postprandial lipid metabolism in healthy subjects. PLoS One, 11, e0160396. Benchekroun, M., Romero, A., Egea, J., Leon, R., Michalska, P., Buendia, I., ... Ismaili, L. (2016). The antioxidant additive approach for Alzheimer's disease therapy: new ferulic (Lipoic) acid plus melatonin modified tacrines as cholinesterases inhibitors, direct antioxidants, and nuclear factor (erythroid-derived 2)-like 2 activators. Journal of Medicinal Chemistry, 59, 9967–9973. Carden, T. J., Hang, J., Dussault, P. H., & Carr, T. P. (2015). Dietary plant sterol esters must be hydrolyzed to reduce intestinal cholesterol absorption in hamsters. Journal of Nutrition, 145, 1402–1407. Danesi, F., Gomez-Caravaca, A. M., de Biase, D., Verardo, V., & Bordoni, A. (2016). New insight into the cholesterol-lowering effect of phytosterols in rat cardiomyocytes. Food Research International, 89, 1056–1063. Derosa, G., D'Angelo, A., Romano, D., & Maffioli, P. (2016). A clinical trial about a food supplement containing alpha-lipoic acid on oxidative stress markers in type 2 diabetic patients. International Journal of Molecular Sciences, 17, 1802. Dong, S., Zhang, R., Ji, Y. C., Hao, J. Y., Ma, W. W., Chen, X. D., ... Yu, H. L. (2016). Soy milk powder supplemented with phytosterol esters reduced serum cholesterol level in hypercholesterolemia independently of lipoprotein E genotype: a random clinical placebo-controlled trial. Nutrition Research, 36, 879–884. Fu, Y., Zhang, Y., Hu, H., Chen, Y., Wang, R., Li, D., & Liu, S. (2014). Design and straightforward synthesis of novel galloyl phytosterols with excellent antioxidant activity. Food Chemistry, 163, 171–177. Gu, S., Wang, J., Wei, X., Cui, H., Wu, X., & Wu, F. (2014). Enhancement of lipasecatalyzed synthesis of caffeic acid phenethyl ester in ionic liquid with DMSO cosolvent. Chinese Journal of Chemical Engineering, 22, 1314–1321. He, W. S., Hu, D., Wang, Y., Chen, X. Y., Jia, C. S., Ma, H. L., & Feng, B. (2016). A novel chemo-enzymatic synthesis of hydrophilic phytosterol derivatives. Food Chemistry, 192, 557–565. He, W. S., Jia, C. S., Ma, Y., Yang, Y. B., Zhang, X. M., Feng, B., & Yue, L. (2010). Lipasecatalyzed synthesis of phytostanyl esters in non-aqueous media. Journal of Molecular Catalysis B: Enzymatic, 67, 60–65. He, W. S., Ma, Y., Pan, X. X., Li, J. J., Wang, M. G., Yang, Y. B., ... Feng, B. (2012). Efficient solvent-free synthesis of phytostanyl esters in the presence of acid-
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Food Chemistry journal homepage: www.elsevier.com/locate/foodchem
Enzymatic synthesis of phytosteryl lipoate and its antioxidant properties ⁎
MARK
Huiqi Wang, Chengsheng Jia , Xue Xia, Eric Karangwa, Xiaoming Zhang State Key Laboratory of Food Science and Technology, School of Food Science and Technology, Jiangnan University, Wuxi 214122, Jiangsu, PR China
A R T I C L E I N F O
A B S T R A C T
Chemical compounds studied in this article include: Phytosterols (PubChem CID: 12303662) Lipoic acid (PubChem CID: 6112) Stigmasterol (PubChem CID: 5280794)
In this work, an enzymatic route for synthesizing phytosteryl lipoate was successfully set up for the first time. The structure of final product phytosteryl lipoate was determined by Fourier Transform Infrared (FTIR), Mass Spectrometry (MS) and Nuclear Magnetic Resonance (NMR). The maximum conversion of 71.2% was achieved when the following conditions were employed: 150 mmol/L phytosterol, 1: 2.5 M ratio of phytosterol to lipoic acid, 10 g/L of 4 Å molecular sieves and 60 g/L Candida rugosa in 2-methyl-2-butanol/n-hexane (1:1, v/v) at 55 °C for 96 h. The physicochemical properties including solubility and antioxidant ability of phytosteryl lipoate in oil were assessed. The results revealed that phytosteryl lipoate possessed over twice as much oil solubility as free phytosterol and also showed better antioxidant ability. Investigation on its biological functions will be the main object in the future work.
Keywords: Phytosteryl lipoate Enzymatic synthesis Oil solubility Peroxide value Antioxidant activity
1. Introduction Phytosterol plays important roles in pharmaceutical industries because of the vast physiological significances such as cholesterol-lowering, antioxidant and anticancer functions (Alappat, Valerio, & Awad, 2010; Bard, Paillard, & Lecerf, 2015; Danesi, Gomez-Caravaca, de Biase, Verardo, & Bordoni, 2016; Woyengo, Ramprasath, & Jones, 2009). In addition, plant sterols are one of healthy factors in food (Baumgartner, Mensink, & Plat, 2016; Dong et al., 2016; te Velde et al., 2015). However, the practical application of phytosterol is limited due to its poor solubility in both water and oil, which is caused by its special chemical structure. Consequently, there is a need to modify phytosterol structure with its functions retained. Up to now, most studies have focused on the esterification of phytosterol with fatty acids to improve its solubility, however, there are a few reports on phytosteryl esters with their improved biological functions (Fu et al., 2014; Tan & Shahidi, 2012, 2013). What is noteworthy is that these phytosteryl esters can be decomposed into two initial reactants after digestion (Carden, Hang, Dussault, & Carr, 2015; Lubinus, Barnsteiner, Skurk, Hauner, & Engel, 2013). As a result, it’s crucial to choose a suitable reactant which can improve respectively the physicochemical properties and can also perform their beneficial functions together. Antioxidants have been always attracting the attention of many researchers, since they can prevent not only the oxidation of fat rich food to prolong their storage time but also senescence and some other diseases caused by reactive oxygen species (ROS) and free radicals in
⁎
Corresponding author. E-mail address: [email protected] (C. Jia).
http://dx.doi.org/10.1016/j.foodchem.2017.08.025 Received 31 May 2017; Received in revised form 30 July 2017; Accepted 4 August 2017 Available online 04 August 2017 0308-8146/ © 2017 Published by Elsevier Ltd.
body, such as Alzheimer’s disease (Rosini et al., 2005). Lipoic acid is a very important kind of vitamin B compounds and mainly presents in the liver tissue of animals and some plants such as tomato, broccoli, carrot and spinach. The high electron density in closed five-membered ring imparts lipoic acid significant electrophilic property and the ability to react with free radicals. In the body, lipoic acid can be reduced into dihydrogen lipoic acid which has two sulfydryls. Both lipoic acid and dihydrogen lipoic acid have good resistance to oxidation and can work together in fat soluble and water soluble environment. In this relation, they are widely known as natural universal antioxidants. Lipoic acid owns various useful functions for human health, which has been already used in medical treatment, such as the prevention of diabetes, cancer, cardiovascular disease and inflammation (Benchekroun et al., 2016; Derosa, D'Angelo, Romano, & Maffioli, 2016; Moura, de Andrade, dos Santos, & Goulart, 2015; Tsuji-Naito, Ishikura, Akagawa, & Saeki, 2010; Ying et al., 2010). Meanwhile, it is also extensively applied in cosmetics and health food fields (Leysen & Aerts, 2016; Merry, Kirk, & Goyns, 2008). Researchers have already reported the synthesis of lipoic acid conjugates for health applications (Hsieh et al., 2015; Kaki, Balakrishna, & Prasad, 2014; Lahiani, Hidmi, Katzhendler, Yavin, & Lazarovici, 2016). Recent study has demonstrated that the phytosterol/lipoic acid combination could work together and played a preferable and superior cholesterol-lowering role than phytosterol or lipoic acid alone (Rideout et al., 2016). Phytosteryl lipoate as shown in Fig. 1, synthesized through esterification of phytosterol with lipoic acid, was supposed to play better physiological functions with smaller
Food Chemistry 240 (2018) 736–742
H. Wang et al. O
Fig. 1. Synthetic route of phytosteryl lipoate. OH
R
S S
Lipoic acid
O
Lipase
+
O S S
R
R=
,
,
,
etc
Phytosteryl lipoate
HO R=
,
,
,
etc
Phytosterol
2.3. Purification of esterified reaction mixture
amounts than either phytosterol or lipoic acid. Enzymatic method is a great route for mild reaction, high safety and less by-products, which meets the requirements for good health and green. In recent years, lipases have been widely used for catalyzing esterification reactions of phytosterols. Schär and Nyström produced steryl ferulates using enzymatic method for the first time (Schär & Nyström, 2016). Miao et al. have selected Candida rugosa as catalyst to develop phytosteryl laurate in non-aqueous media (Miao et al., 2014). To the best of our knowledge, the enzymatic synthesis of phytosteryl lipoate has not still been well documented. The present work was therefore aimed to explore a lipase-catalyzed method to synthesize phytosteryl lipoate and investigate its optimal reaction conditions. After purification by Thin-Layer Chromatography (TLC) and silica gel column chromatography, final product’s chemical structure was determined by FT-IR, NMR and MS. Meanwhile, the solubility and antioxidant activity of phytosteryl lipoate were also evaluated.
After the esterification reaction, the mixture was filtered to remove the lipase and molecular sieves, and then the extract was treated with 0.1 M NaHCO3 aqueous solution to neutralize excess lipoic acid. The filtrates were evaporated by rotary evaporator to remove the solvents, then, 1.0 g of the dried sample was dissolved in petroleum ether (60–90 °C)/ethyl acetate/formic acid mixture (15:1:0.02, v/v/v). The obtained solutions were subsequently applied to a silica gel column (24 × 1200 mm) and eluted with petroleum ether (60–90 °C)/ethyl acetate/formic acid (15:1:0.02, v/v/v) at a flow rate of 0.5 mL/min. The eluate was detected by HPLC and collected at the same time, the esters were then obtained after the solvents evaporation by rotary evaporation. 2.4. HPLC analysis The samples (10 µL) taken out from reaction systems were injected in Waters 1525 HPLC analysis equipped with a symmetry-C18 column (5 μm, 4.6 × 150 mm, Waters, USA) at 45 °C. The sample was eluted with methanol (mobile phase) at a flow rate of 1 mL/min. The effluent was monitored with a Grace Alltech 3300 evaporative light scattering detector (ELSD) at 55 °C and nitrogen was used as the carrier gas at a flow rate of 1.5 L/min. The esterification rate of phytosterol with lipoic acid to form phytosteryl lipoate was calculated via calibration curve and purified phytosteryl lipoate was used as external standard. The conversion was defined as follows:
2. Materials and methods 2.1. Chemicals Phytosterols were a generous gift from Jiangsu Spring Fruit Biological Products Co., Ltd. (Taixing, P.R. China). The purity of plant sterols was > 97% (63% β-sitosterol and 37% stigmasterol). Lipoic acid was purchased from Xi’an Jinheng Chemical Co., Ltd. (Shanxi, China). Novozym 435 (lipase B from Candida antarctica, immobilized on a macroporous acrylic resin, 10,000 PLU/g), Lipozyme TL IM (lipase from Thermomyces lanuginosus, immobilized on silica granulation, 250 IUN/g), Lipozyme RM IM (lipase from Rhizomucor miehei, immobilized on an anionic exchange resin, 275 IUN/g) were obtained from Novo Nordisk Co., Ltd. (Shanghai, China). Candida rugosa (lyophilized powder, Type VII, 847 U/mg) was supplied by Sigma-Aldrich Co., Ltd. (Shanghai, China). Methanol of spectral grade was obtained from Tedia Company Inc. (Shanghai, China). The rapeseed oil was purchased from the local supermarket. The 3 Å and 4 Å molecular sieves and other common reagents of analytical grade were from Sinopharm Chemical Reagent Co., Ltd. (Shanghai, China).
XPSE (%) = CPSE /CPS × 100
(1)
where CPSE is the actual concentration of phytosteryl lipoate at the end of the reaction, CPS is the initial concentration of phytosterol. 2.5. FTIR analysis The purified phytosteryl lipoate was dried completely under vacuum and then mixed with KBr. FTIR analysis was conducted on a FT-IR spectrophotometer (Thermo Nicolet iS10 FTIR, Thermo Electron, USA) using an attenuated total reflectance method with the spectral scanning scope for 400–4000 cm−1, the number of scans is 32 and resolution is 4 cm−1.
2.2. Enzymatic synthesis of phytosteryl lipoate
2.6. MS analysis
Typically, phytosterol (1 mmol) and lipoic acid (1–4 mmol) were dissolved in 10 mL organic solvent at first, followed by the addition of 0–0.4 g of molecular sieves and lipase (0.4–0.6 g). The reaction was carried in a shaking water bath at 150 r/min, for 48–120 h at 40–60 °C. All samples were performed in triplicate.
Liquid chromatography mass spectrometry (Maldi Synapt Q-Tof, Waters, USA) was used to further identify the stigmasteryl ester under positive-ion electron spray ionization (ESI) mode. The MS parameters were listed as follows: capillary voltage, 3.5 kV; cone voltage, 30 V; source block temperature, 100 °C; desolvation temperature, 400 °C; 737
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3. Results and discussion 3.1. Structural analysis of phytosteryl lipoate 3.1.1. HPLC analysis The conversion and purity of the final product were determined by HPLC. At the end of the reaction, the reaction mixture was quantitatively analyzed by HPLC and the results were presented in Fig. S1. The peaks of phytosterol and lipoic acid were confirmed by their respective standards. The phytosterol used in the present experiment mainly contained stigmasterol and sitosterol, consequently, the synthetic phytosteryl esters were stigmasteryl lipoate and sitosteryl lipoate. According to the spectrogram and polarity analysis, lipoic acid was firstly eluted at the retention time of 1.745 min, and phytosterols (stigmasterol and sitosterol) were then eluted at the retention time of 3.128 and 3.374 min, respectively. Additionally, two new contiguous peaks at 6.064 min and 6.632 min appeared and these compounds might be stigmasteryl lipoate and sitosteryl lipoate, respectively. Fig. 2. FTIR spectra of lipoic acid (a) and phytosteryl lipoate (b).
3.1.2. FTIR analysis The IR spectra of lipoic acid and the newly synthesized product were compared, as shown in Fig. 2(a), the broad peak between 2400 cm−1 and 3300 cm−1 indicated the presence of eOH group from eCOOH. The strong peak at 1692.84 cm−1 was the signal of the C]O group from eCOOH. In Fig. 2(b), the peaks at 2898.38 cm−1 and 1366.80 cm−1 were the stretching vibration and bending vibrations of CeH groups for methylene groups, respectively. The peaks at 2866.94 cm−1 and 1462.24 cm−1 were the stretching vibration and bending vibrations of CeH groups for methyl groups. The strong peak at 1732.46 cm−1 was the signal of the C]O group, and the medium peaks at 1173.43 cm−1 and 1131.68 cm−1 were the stretching vibration of CeO group of ester bond. In summary, the disappeared absorption signal of free carboxyl group and the emerging ester bond provided evidence that the new product was phytosteryl lipoate.
desolvation gas flow, 700 lit/hr; cone gas flow rate, 50 l/h and scan range, m/z 50–1000. 2.7. NMR analysis NMR experiments were performed on an NMR spectrometer (Avance III 400 MHz, Bruker, Switzerland) using CDCl3 as the solvent and tetramethylsilane as the internal standard. The 1H and 13C NMR spectrum were acquired at 400.14 and 100.62 MHz at 298.0 K, respectively. 2.8. Determination of oil solubility Due to the complex oil components, the solubility of phytosterol and phytosteryl lipoate was determined by direct observation, and phytosteryl ferulate was also measured as a reference. The method was described as follows: 200 mg samples were put into a flask while stirred and heated to 30 °C, then rapeseed oil was added drop by drop until the all samples were entirely dissolved and the oil amount of consumption was noted down immediately. The oil solubility was represented as grams (g) per litre (L) at 30 °C.
3.1.3. MS analysis HPLC-MS with ESI was used to further confirm stigmasteryl lipoate under positive-ion mode. The main advantage of this method is that the reaction mixture could be used for analysis without separation and purification prior to analysis (He et al., 2014). Theoretically, the relative molecular weight for stigmasteryl lipoate was 600. As shown in Fig. 3, the molecular ion [M+H]+ at m/z 601.4 and a sodium adduct molecular ion [M+Na]+ at m/z 623.4 were also identified. The molecular ions [M+H]+ and [M+Na]+ were also observed in previous reports about phytosteryl esters using ESI-MS with positive-ion mode (He et al., 2012, 2016). In addition, a major fragment [M+H-lipoic acid]+ at m/z 395.4 was observed. Similar [M+H-ferulic acid]+ and [M+H-lauric acid]+ fragments have been also reported in the MS analysis of campesteryl ferulate and ergosterol laurate (He et al., 2014; Tan & Shahidi, 2011). These results proved that the synthesized product was stigmasteryl lipoate.
2.9. Antioxidant activity studies in oil The antioxidant ability of phytosteryl lipoate (purity, > 98%) in edible oil was measured by peroxide value (POV) method. It is wellknown that edible oils are full of various unsaturated fatty acids which are subjected to oxidation easily (Köckritz & Martin, 2008). The edible oils containing samples firstly dissolved in chloroform and glacial acetic acid, then the peroxide in the oils would generate iodine after reacting with potassium iodide, finally the amount of iodine was measured using the standard solution of sodium thiosulfate by titration and POV can be indirectly calculated. The specific method was described by Min et al. (2014) with slight modification according to GB/T5009.227-2016, and the POV was calculated using the following equation:
X1 = (V−V0) × C× 0.1269 × 100/m
3.1.4. NMR analysis The synthesized and purified stigmasteryl lipoate was analyzed by 1 H and 13C NMR. The results were listed as follows: 1H NMR (400 MHz, CDCl3): δ = 0.63 (3 H, s, 18-H), 0.73 (6 H, d, J = 7.2 Hz, 26–27-H), 0.78 (3 H, d, J = 6.4 Hz, 21-H), 0.89–0.98 (6 H, m), 0.95 (3 H, s, 19-H), 1.01–1.23 (6 H, m), 1.32–1.66 (16 H, m, 3′-H, 4′-H, 5′-H, etc.), 1.77–2.08 (5 H, m, 7′-Ha, etc.), 2.21–2.25 (4 H, m, 2′-H, etc.), 2.36–2.43 (1 H, dt, J = 4.0, 8.0 Hz, 7′-Hb), 3.02–3.15 (2 H, m, 8′-H), 3.47–3.54 (1 H, m, 6′-H), 4.52–4.58 (1 H, m, 3-H), 4.94 (1 H, dd, J = 8.0, 12.0 Hz, 22-H or 23-H), 5.08 (1 H, dd, J = 8.0, 12.0 Hz, 22-H or 23-H), 5.30 (1 H, d, J = 4.0, 6-H). 13C NMR (100 MHz, CDCl3): δ = 12.06 (29CH3), 12.26 (18-CH3), 18.99 (21-CH3), 19.34 (19-CH3), 21.03 (26CH3), 21.10 (27-CH3), 21.23 (CH2), 24.37 (11-CH2), 24.78 (3′-CH2),
(2)
The X1 represented POV (g/100 g), V was the volume (mL) of standard solution of sodium thiosulfate consumed by samples, while the volume of the sample blank was expressed as V0 (mL). C represented the concentration of sodium thiosulfate (0.002 mol/L), and m (g) corresponded to the mass of the samples. The number 0.1269 was the mass of iodine which was equal to 1.00 mL standard solution of sodium thiosulfate (Na2S2O3 = 1.00 mol/L) and 100 was the conversion coefficient. 738
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Fig. 3. HPLC-MS chromatogram of stigmasteryl lipoate.
25.42 (15-CH2), 27.83 (CH2), 28.74 (4′-CH2), 28.92 (CH2), 31.87 (8- or 25-CH), 31.89 (8- or 25-CH), 34.45 (5′-CH2), 34.61 (8′-CH2), 36.63 (10C), 37.01 (CH2), 38.17 (CH2), 38.49 (CH2), 39.64 (CH2), 40.23 (20-CH), 40.50 (7′-CH), 42.22 (13-C), 50.06 (9-CH), 51.24 (24-CH), 55.95 (17CH), 56.37 (6′-CH), 56.79 (14-CH), 73.87 (3-C), 122.63 (6-C), 129.34 (22-C), 138.31 (23-C), 139.72 (5-C), 172.91 (C = O). The 1H NMR and 13 C NMR spectra of phytosteryl lipoate were displayed in Figs. S2 and S3, respectively. The proton signals were overlapped at δ 0.63–3.15 in 1H NMR, the chemical shift below δ 1.2 could primarily be attributed to methyl groups in stigmasterol and those between δ 1.2 and 3.2 were ascribed to methylene in both stigmasterol and lipoic acid. The proton peaks of lipoic acid in the product were at δ 1.32–1.66, 1.77–2.08, 2.21–2.25, 2.36–2.43, 3.02–3.15 and 3.47–3.54, respectively. These results are greatly consistent with the findings reported by Madawala, Andersson, Jastrebova, Almeida, and Dutta (2012). The chemical shift of 3-position proton was at δ 3.5 in free phytosterol (He et al., 2016), while it shifted to δ 4.5 in the product due to the formation of ester bond, which agreed with its resonances in other phytosteryl esters (Kobayashi et al., 2014). Similarly, the 13C NMR analysis of the compound showed that the chemical shift of 3-C (δ 73.87) was lower than that in free stigmasterol (δ 71.8) (Huang, Cao, Xu, & Chen, 2011).
Table 1 Influence of different solvents and lipases on conversion. Solvent
tert-Butanol 2-Methyl-2-butanol n-Hexane Acetone/n-hexane (1:1, v/v) 2-Methyl-2-butanol/nhexane (1:1, v/v) tert-Butanol/n-hexane (1:1, v/v)
Conversion (%) Novozyme 435
Candida Rugosa
TL IM
RM IM
0.8 0.6 0.2 3.2 1.3
0.6 1.7 0.4 5.3 6.5
0.4 0.6 0.2 1.7 3.4
0.2 0.3 0.1 0.2 0.1
0.8
1.2
2.2
0.6
Reaction condition: 50 mmol/L phytosterol, 1.5:1 M ratio of lipoic acid to phytosterol, 20 g/L lipase and 40 g/L of 4 Å molecular sieve at 150 r/min and 45 °C for 72 h.
shown in Table 1. This could be attributed to the poor solubility of lipoic acid in n-hexane. On the contrary, though both substrates have good solubility in tert-butanol and 2-methyl-2-butanol, the deprivation of water made the enzyme inactive, hence the conversion was low. In such situation, mixed solvents are often used when the significant difference exists between the polarities of two substrates (Gu et al., 2014; Zhao et al., 2011). Thus, n-hexane was mixed with the three solvents of different polarities (1:1, v/v), respectively, and the results showed that the conversion was significantly improved in co-solvent system, especially in acetone/n-hexane and 2-methyl-2-butanol/n-hexane system, respectively. Four lipases in different forms were used in the present study. Novozyme 435, Lipozyme TL IM and Lipozyme RM IM are all immobilized lipases, while Candida rugosa are powder lipase. Different lipases, originated from diverse sources of microorganism, presented dramatically different catalytic efficiency for the esterification of phytosterol with lipoic acid. The catalytic efficiency of Lipozyme RM IM was the lowest for the reaction conversion. Novozyme 435 in acetone/ n-hexane (1:1, v/v) showed the highest conversion of 3.2% and
3.2. Lipase-catalyzed synthesis of phytosteryl lipoate 3.2.1. Screen of enzyme and solvent Different solvents, with log P values from −0.26 to 3.50, and several kinds of lipase are screened and their conversions are recorded in Table 1. The influences of solvent are mainly embodied in two aspects: on the one hand, the solubility of substrates is a pivotal factor for esterification rate, on the other hand, every enzyme needs a certain amount of water to maintain its configuration and activity, and the higher the polarity of solvent is, the lower the activity of enzyme is. nHexane is a commonly used solvent with low polarity and food safety, however, its conversion was the lowest among the selected solvents as 739
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Fig. 4. Influence of phytosterol concentration (a), molar ratio (b), molecular sieve (c), lipase load (d), temperature (e) and time (f) on esterification rate. (Reaction condition: (a) 1.5:1 M ratio of lipoic acid to phytosterol, 20 g/L lipase and 40 g/L of 4 Å molecular sieve, 45 °C, 72 h; (b) 150 mmol/L phytosterol, 20 g/L lipase and 40 g/L of 4 Å molecular sieve, 45 °C, 72 h; (c) 150 mmol/L phytosterol, 2.5:1 M ratio of lipoic acid to phytosterol and 20 g/L lipase, 45 °C, 72 h; (d) 150 mmol/L phytosterol, 2.5:1 M ratio of lipoic acid to phytosterol and 10 g/L of 4 Å molecular sieve, 45 °C, 72 h; (e) 150 mmol/L phytosterol, 2.5:1 M ratio of lipoic acid to phytosterol, 60 g/L lipase and 10 g/L of 4 Å molecular sieve, 72 h; (f) 150 mmol/L phytosterol, 2.5:1 M ratio of lipoic acid to phytosterol, 60 g/L lipase and 10 g/L of 4 Å molecular sieve, 55 °C.)
substrates and catalysis temperature used in this study. Kim and Akoh (2007) reported that Candida rugosa was a substrate-dependent enzyme for the synthesis of phytosterol esters. Hence, Candida rugosa in 2-methyl-2-butanol/n-hexane (1:1, v/v) was considered as the most desirable biocatalyst and solvent.
Lipozyme TL IM in 2-methyl-2-butanol/n-hexane (1:1, v/v) showed the maximum conversion of 3.4%. However, Candida rugosa in acetone/nhexane (1:1, v/v) and 2-methyl-2-butanol/n-hexane (1:1, v/v) showed the highest conversion of 5.3% and 6.5%, respectively. It was clear that Candida rugosa showed high catalytic ability compared to the other three immobilized enzymes at 45 °C. Though Novozyme 435 was regarded as the most active lipase in most cases, it could not achieve better conversion than C. Rugosa, which might be ascribed to the
3.2.2. Effect of substrate concentration The influence of different 740
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3.2.7. Effect of reaction time As shown in Fig. 4(f), the esterification rate linearly increased with increasing reaction time from 24 h to 96 h, and the maximum conversion (71.2%) was reached after 96 h. When the reaction prolonged up to 120 h, the esterification rate decreased to 68.9%. Therefore, the reaction had reached equilibrium after around 96 h. According to the above optimization results, the maximum yield (> 71%) could be reached under the selected conditions: 150 mmol/L phytosterol, 1:2.5 M ratio of phytosterol to lipoic acid, 60 g/L Candida rugosa, 10 g/L 4Å molecular sieves in 2-methyl-2-butanol/n-hexane (1:1, v/v) at 55 °C for 96 h at a stirring rate of 150 r/min.
(25–200 mmol/L) on the esterification of phytosterol with lipoic acid was investigated and the results are shown in Fig. 4(a). At first, the conversion linearly increased with increasing substrate concentration, and the conversion reached the highest rate (10.2%) at 150 mol/L phytosterol concentration, then decreased afterward with further increasing concentration. The conversion dropped to 7.8%, this observation might be attributed to the restricted total solubility of substrates and reduced enzyme activity, because enzyme active sites were occupied by redundant substrates (He et al., 2010). 3.2.3. Effect of molar ratio Different molar ratios varied from 1:1.5 to 3:1 of lipoic acid to phytosterol were used in a series experiments. As presented in Fig. 4(b), it has been demonstrated that the excess of either phytosterol or lipoic acid was beneficial for esterification reaction, the conversion was lower (5.6%) at equimolar substrates. Our findings are in good agreement with previous studies (He et al., 2016; No, Zhao, Lee, Lee, & Kim, 2013). Considering the purification procedure, lipoic acid is more convenient to separate than phytosterol, therefore excess lipoic acid was used. As the molar ratio increased from 1:1 to 2.5:1, the esterification rate was boosted from 5.6% to 16.1%, then dropped to 11.3% when the molar ratio was 3:1. This might be attributed to the denaturation of enzyme in acidic condition.
3.3. Comparison of solubility In general, substances can become more hydrophobic with the increase of lipophilic group in the molecules, and their solubility in oils will accordingly be enhanced. The solubility of phytosteryl lipoate and phytosteryl ferulate reached 20.4 g/L and 22.5 g/L in rapeseed oil at 30 °C, respectively, while the solubility of phytosterols was only 9.1 g/L under the same condition. Conclusively, the esterification of phytosterol with lipoic acid improved its solubility in edible oils. 3.4. Antioxidant ability
3.2.4. Effect of molecular sieve concentration Controlling the water content in the reaction system is of great concern. Too much water could make reaction equilibrium move to the left side, while lack of water would have impact on the conformation and the catalytic efficiency of enzyme, then conversion would decline under any of the two situations. Hence, it is very critical to choose appropriate variety and quantity of dehydrating agent. Due to the economic factors and separation procedure, different dosage of 3 Å and 4 Å molecular sieves were used in our study. 3 Å molecular sieve has better ability to remove water compared with 4 Å molecular sieve (He et al., 2010). From Fig. 4(c), it could be concluded that molecular sieve possessed a large effect on esterification rate and the conversion using 4 Å molecular sieve was higher than that using 3 Å molecular sieve. The conversion reached the maximum value (30.2%) when the 4 Å molecular sieve concentration was 10 g/L and drastically declined afterward, while the conversion kept decreasing when 3 Å molecular sieve was used. The water in reaction system was gradually reduced with the increase concentration of molecular sieve, which led to the decline of enzyme activity. Therefore, 10 g/L of 4 Å molecular sieve was selected for further experiments.
The antioxidant ability of phytosteryl lipoate was evaluated by POV determination and was compared to that of phytosterol. Phytosteryl ferulate, the main ingredient of γ-oryzanol which has already been used commercially for its cholesterol-lowering, antioxidant and antiinflammation properties, was selected for reference. All samples were analyzed in triplicate. The same amount (0.2 g/kg) of different samples were added into rapeseed oil and their corresponding POVs were shown in Fig. 5. Since oil samples were continuously heated at 60 °C, all samples would generate more and more peroxide and their POVs increased. According to GB2716-2005, POV should be below 0.25 g/ 100 g in oil. As denoted in Fig. 5, the rapeseed oil without any additives showed the highest POVs, while the POVs of oil samples containing phytosteryl lipoate showed the lowest POVs from 24 h to 84 h. The POVs of blank sample were 0.23 g/100 g at 60 h and 0.27 g/100 g at 72 h. The samples treated with phytosterol had slightly lower POVs, and its POVs were 0.21 g/100 g at 60 h and 0.25 g/100 g at 72 h. It could be seen that both phytosteryl lipoate and phytosteryl ferulate had better oxidation resistance than phytosterol, their POVs were 0.19 g/ 100 g and 0.22 g/100 g at 72 h, 0.23 g/100 g and 0.24 g/100 g at 84 h, respectively. The POVs of phytosteryl lipoate was lower than that of phytosteryl ferulate as a whole, which indicates that phytosteryl lipoate could potentially be used as an antioxidant in oils based food.
3.2.5. Effect of lipase load Fig. 4(d) illustrated the effect of different concentration of Candida rugosa varying from 20 g/L to 80 g/L on the conversion. As displayed in Fig. 4(d), the conversion increased from 28.9% to 38.4% with increasing Candida rugosa load from 20 g/L to 40 g/L, then reached to 47.2% when the lipase loading was 60 g/L. As the loading continued to increase up to 80 g/L, the esterification rate was slightly changed. Out of the economic aspects, 60 g/L lipase loading was selected for the further studies. 3.2.6. Effect of reaction temperature The substrates solubility and enzyme activity are both strongly dependent on the reaction temperature. The effect of reaction temperature (40 °C to 60 °C) on esterification rate was studied. As presented in Fig. 4(e), the initial esterification rate was 41.8% at 40 °C, and the conversion gradually increased with increasing temperature and reached the apex (56.2%) at 55 °C, then decreased with further increasing of the reaction temperature from 55 °C to 60 °C. These findings are in accordance with the results reported by Kim and Akoh (2007), which stipulated that Candida rugosa could play its best catalytic role when the reaction temperature was set at 55 °C.
Fig. 5. Peroxide value (POV) (g/100 g) of rapeseed oil treated with phytosterol, phytosteryl lipoate and phytosteryl ferulate at 60 °C.
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4. Conclusion In this study, the phytosteryl lipoate was successfully synthesized by a direct enzymatic approach. The effects of 8 parameters (solvent, lipase, phytosterol concentration, molar ratio, molecular sieve, lipase load, temperature and time) on conversion were evaluated and the highest conversion was 71.2%. After conjugation with lipoic acid, the oil solubility and antioxidant ability of phytosterol have been greatly improved. This novel ester broadens the application of phytosterol in food and pharmaceutical industry and may also possess physiological functions of phytosterol and lipoic acid, such as cholesterol-lowering ability. Acknowledgement This research was financially supported by the Natural Science Foundation of Jiangsu Province (BK20161133) and program of “Collaborative Innovation Center of Food Safety and Quality Control in Jiangsu Province”. . Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.foodchem.2017.08.025. References Alappat, L., Valerio, M., & Awad, A. B. (2010). Effect of vitamin D and beta-sitosterol on immune function of macrophages. International Immunopharmacology, 10, 1390–1396. Bard, J. M., Paillard, F., & Lecerf, J. M. (2015). Effect of phytosterols/stanols on LDL concentration and other surrogate markers of cardiovascular risk. Diabetes Metabolism, 41, 69–75. Baumgartner, S., Mensink, R. P., & Plat, J. (2016). Effects of a plant sterol or stanol enriched mixed meal on postprandial lipid metabolism in healthy subjects. PLoS One, 11, e0160396. Benchekroun, M., Romero, A., Egea, J., Leon, R., Michalska, P., Buendia, I., ... Ismaili, L. (2016). The antioxidant additive approach for Alzheimer's disease therapy: new ferulic (Lipoic) acid plus melatonin modified tacrines as cholinesterases inhibitors, direct antioxidants, and nuclear factor (erythroid-derived 2)-like 2 activators. Journal of Medicinal Chemistry, 59, 9967–9973. Carden, T. J., Hang, J., Dussault, P. H., & Carr, T. P. (2015). Dietary plant sterol esters must be hydrolyzed to reduce intestinal cholesterol absorption in hamsters. Journal of Nutrition, 145, 1402–1407. Danesi, F., Gomez-Caravaca, A. M., de Biase, D., Verardo, V., & Bordoni, A. (2016). New insight into the cholesterol-lowering effect of phytosterols in rat cardiomyocytes. Food Research International, 89, 1056–1063. Derosa, G., D'Angelo, A., Romano, D., & Maffioli, P. (2016). A clinical trial about a food supplement containing alpha-lipoic acid on oxidative stress markers in type 2 diabetic patients. International Journal of Molecular Sciences, 17, 1802. Dong, S., Zhang, R., Ji, Y. C., Hao, J. Y., Ma, W. W., Chen, X. D., ... Yu, H. L. (2016). Soy milk powder supplemented with phytosterol esters reduced serum cholesterol level in hypercholesterolemia independently of lipoprotein E genotype: a random clinical placebo-controlled trial. Nutrition Research, 36, 879–884. Fu, Y., Zhang, Y., Hu, H., Chen, Y., Wang, R., Li, D., & Liu, S. (2014). Design and straightforward synthesis of novel galloyl phytosterols with excellent antioxidant activity. Food Chemistry, 163, 171–177. Gu, S., Wang, J., Wei, X., Cui, H., Wu, X., & Wu, F. (2014). Enhancement of lipasecatalyzed synthesis of caffeic acid phenethyl ester in ionic liquid with DMSO cosolvent. Chinese Journal of Chemical Engineering, 22, 1314–1321. He, W. S., Hu, D., Wang, Y., Chen, X. Y., Jia, C. S., Ma, H. L., & Feng, B. (2016). A novel chemo-enzymatic synthesis of hydrophilic phytosterol derivatives. Food Chemistry, 192, 557–565. He, W. S., Jia, C. S., Ma, Y., Yang, Y. B., Zhang, X. M., Feng, B., & Yue, L. (2010). Lipasecatalyzed synthesis of phytostanyl esters in non-aqueous media. Journal of Molecular Catalysis B: Enzymatic, 67, 60–65. He, W. S., Ma, Y., Pan, X. X., Li, J. J., Wang, M. G., Yang, Y. B., ... Feng, B. (2012). Efficient solvent-free synthesis of phytostanyl esters in the presence of acid-
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