Production of Novel “Functional Oil” Rich in Diglycerides and

Article pubs.acs.org/JAFC

Production of Novel “Functional Oil” Rich in Diglycerides and Phytosterol Esters with “One-Pot” Enzymatic Transesterification Ming-Ming Zheng,∥ Qing Huang,∥ Feng-Hong Huang,* Ping-Mei Guo, Xia Xiang, Qian-Chun Deng, Wen-Lin Li, Chu-Yun Wan, and Chang Zheng Oil Crops Research Institute, Chinese Academy of Agricultural Sciences, Hubei Key Laboratory of Oilcrops Lipid Chemistry and Nutrition, Wuhan 430062, China ABSTRACT: Diglycerides and phytosterol esters are two important functional lipids. Phytosterol esters mixed with dietary diglyceride could not only influence body weight but also prevent or reverse insulin resistance and hyperlipidemia. In this study, a kind of novel “functional oil” rich in both diglycerides and phytosterol esters was prepared with “one-pot” enzymatic transesterification. First, lipase AYS (Candida rugosa) was immobilized on the porous cross-linked polystyrene resin beads (NKA) via hydrophobic interaction. The resulting immobilized AYS showed much better transesterification activity and thermal stability to freeways. On the basis of the excellent biocatalyst prepared, a method for high-efficiency enzymatic esterification of phytosterols with different triglycerides to produce corresponding functional oils rich in both diglycerides and phytosterol esters was developed. Four functional oils rich in both diglycerides and phytosterol esters with conversions >92.1% and controllable fatty acid composition were obtained under the optimized conditions: 80 mmol/L phytosterols, 160 mmol/L triglycerides, and 25 mg/mL AYS@NKA at 180 rpm and 50 °C for 12 h in hexane. The prepared functional oil possessed low acid value (≤1.0 mgKOH/g), peroxide value (≤2.1 mmol/kg), and conjugated diene value (≤1.96 mmol/kg) and high diglyceride and phytosterol ester contents (≥10.4 and ≥20.2%, respectively). All of the characteristics favored the wide application of the functional oil in different fields of functional food. KEYWORDS: diglycerides, phytosterol esters, functional oil, enzymatic transesterification

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INTRODUCTION Phytosterols have attracted great attention because they are known to have a hypocholesterolemic effect by lowering plasma total and low-density lipoprotein (LDL) cholesterol levels without affecting plasma high-density lipoprotein (HDL) cholesterol concentration. Phytosterol esters (PE), derived from phytosterols and inheriting all of the excellent properties of phytosterols mentioned above, have a much greater solubility in oils and a lower melting point as compared to the corresponding phytosterols. These advantages make it easier to incorporate phytosterol esters into a wide variety of diets and fat-based food products. 1−3 Besides phytosterol esters, diglycerides (DG) are also reported as a new kind commendable functional lipid.4−7 The structural and metabolic characteristics of diglycerides compared with triglycerides appear to be responsible for suppression of body fat accumulation, body weight loss, and lower postprandial serum triglyceride levels.8−10 Recently, Ehud et al. pointed out that phytosterol esters mixed with dietary diglyceride could not only influence body weight but also prevent or reverse insulin resistance and hyperlipidemia; thus, they could be serve as functional ingredients for metabolic syndrome or diabetic sufferers.11 Because phytosterol esters and diglycerides scarcely exist in nature, there is an urgent need to produce them with an efficient and economical process. Diglycerides and phytosterol esters can be synthesized by either chemical or enzymatic processes.12,13 However, besides the high reaction temperature and lack of selectivity, the chemical methods involve many other disadvantages including inevitable side products, high © 2014 American Chemical Society

energy consumption, poor product quality, and low yield. Enzymatic catalysis, which proceeds efficiently under mild conditions, produces fewer byproducts, and benefits from the biocatalyst’s high specificity, is attractive for the synthesis of phytosterol esters and diglycerides. Compared to the current chemical methodologies, it provides for an environmentally friendlier, more energy efficient, and potentially more costeffective techniques due to lowenergy demand and easier downstream processing.14,15 Generally speaking, phytosterol esters and diglycerides have to be prepared separately. Lipasecatalyzed synthesis of phytosterol esters as well as diglycerides has been reported and is attracted more and more attention.16−19 In our previous study, enzymatic synthesis of phytosterol esters was developed with both fatty acid and triglycerides as the acyl donors in high yield.20 However, little attention has been paid to the detailed constitution of the complicated products obtained in phytosterol esterification with triglycerides. According to our later research, the products of our previous study are rich in both phytosterol esters and diglycerides, which make them promising for the design and development of novel functional oils with fat-soluble bioactive ingredients. One-pot synthesis is a strategy whereby a reactant is subjected to successive chemical reactions in just one reactor. It could avoid a lengthy separation process, and purification of Received: Revised: Accepted: Published: 5142

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Figure 1. Preparation scheme of functional oil rich in both phytosterol esters and diglycerides using AYS@NKA as the biocatalyst. beads via adsorption was studied in phosphate buffer (400 mL, 100 mM, pH 7.0). The initial concentration of lipase was kept at 0.5−7.0 mg/mL in buffer solution. The immobilization experiment was conducted at 30 °C for 8 h with continuous stirring. After immobilization, the NKA beads were separated from the lipase solution and then washed with buffer solution three times and then washed with acetone twice. The resulting immobilized lipase (designated AYS@NKA) was lyophilized and stored at 4 °C prior to use. The amount of immobilized lipase was obtained by using the equation

the intermediate chemical compounds would save time and resources.21 The main objective of this study is to develop a novel functional oil rich in both phytosterol esters and diglycerides with one-pot enzymatic esterification. Lipase immobilized on porous cross-linked polystyrene resin beads (NKA) was prepared, characterized, and used as the biocatalyst. The transesterification activity of immobilized lipase was compared with that of the free lipase. Then, a rapid and convenient method was proposed for the enzymatic transesterification of phytosterols with different vegetable oils to produce functional oils in one pot. The physiochemical properties of four kinds of functional oils with phytosterol esters and diglycerides as well as the different fatty acid (FA) compositions were also determined. As far as we are aware, a method related to producing functional oils containing both phytosterol esters and diglycerides with controllable FA composition has not been reported yet.

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Q = [(C0 − C)V ]/M

(1)

where Q is the amount of lipase immobilized onto NKA (mg/g), C0 and C are the concentrations of the lipase in the solutions before and after immobilization, respectively (mg/mL), V is the volume of the aqueous solution (mL), and M is the mass of the beads (g). The amounts of protein in the medium and wash solutions were determined according to the Bradford method.23 Nitrogen sorption experiments were carried out at 77 K using Gemini V2380 surface area and pore size analyzer (Micromeritics, Norcross, GA, USA). One-Pot Enzymatic Synthesis of Functional Oil. The following esterification conditions were used: phytosterols (50−150 mmol/L), triglycerides such as sunflower oil, corn oil, rapeseed oil, or linseed oil (80−640 mmol/L), lipase (free AYS, AYS@NKA, 10−40 mg/mL), and solvent (hexane, iso-octane, etc., 10 mL) were added into an Erlenmeyer flask. The solvent had, in advance, been dehydrated with 15% (w/w) molecular sieves 3 Å for at least 24 h. The vial was placed in a shaking incubator at 45−60 °C with a shaking speed of 180 rpm for a certain time. The reaction bioconversion was monitored periodically by HPLC to confirm production. The bioconversion was calculated from the ratio between the measured concentration of phytosterol esters and the expected concentration assuming complete reaction. Standard solution calibration curves were used to quantify target analytes, as it was found that no matrix effect was present. Calibration curves for phytosterol esters and diglycerides were obtained by dissolving standard substances with mobile phase in different concentrations, and results were detected and recorded on the HPLC-PDA detection. The reaction lasted for a certain time, and the relevant samples (50 μL) were withdrawn from the reaction mixture at relevant time intervals and diluted to 1 mL with the mobile phase for the analytical monitoring of the reaction. The transesterification of phytosterols might occur at either sn-1,3 or sn-2 position of triglyceride; thus, the products including diglycerides and phytosterol esters might be several possibilities (see Figure 1). Following the lipase-catalyzed esterification, immobilized lipase and molecular sieves were filtered from the mixture. Functional oil was prepared after rotary evaporation to remove reaction solvent. Isolation and Analysis of Functional Ingredients in the Functional Oil. High-performance liquid chromatography (HPLC) analysis was performed on an LC-6A Prominence instrument (Shimadzu, Japan) equipped with a photodiode array (PDA) detector. An Inertsil ODS-SP column (5 μm, 250 mm × 4.6 mm) purchased

EXPERIMENTAL PROCEDURES

Reagents and Chemicals. The porous cross-linked polystyrene resin beads (NKA, 200−220 Å) used for preparing the immobilized lipase were obtained from the Chemical Plant of NanKai University (Tianjin, China). Ethanol, 2-propanol, acetonitrile, iso-octane, nhexane, tert-amyl alcohol, cyclohexane, anhydrous disodium phosphate, molecular sieves (3 Å), and other solvents were all purchased from Sinopharm Chemical Reagent (Shanghai, China). 2-Propanol and acetonitrile were of chromatographic pure grade, and other reagents were of analytical reagent grade. Purified water was obtained with a Millipore water purification equipment (Boston, MA, USA). Thin layer chromatography (TLC) silica gel 60 glass plates were purchased from Merck (Darmstadt, Germany). Lipase PS (Burkholderia cepacia lyophilized powder) and AYS (Candida rugsa, lyophilized powder) was purchased from Amano Enzyme Inc. (Tokyo, Japan). Novozym 435 (Candida antarctica), Lipozyme TL IM (Thermomyces lanuginosus), and Lipozyme RM IM (Rhizomucor miehei) were purchased from Novozymes A/S (Bagsværd, Denmark). Phytosterols (β-sitosterol (89.0%), campesterol (8.9%), stigmasterol (2.1%)) were purchased from Vita-Solar Biotechnology Co., Ltd. (Xi’an, China). Refined and bleached sunflower oil, rapeseed oil, corn oil, and linseed oil, which were used as the source of triglycerides, were purchased from the supermarket. Diglyceride and monoglyceride standards were purchased from Larodan Fine Chemicals (Malmö, Sweden). Phytosterol ester standard (97%) was purchased from BASF (China) Co., Ltd. (Shanghai, China). Immobilization of Lipase AYS on the NKA Beads. The immobilized lipase AYS was prepared as proposed by Yan et al. with some modification.22 First, 20.0 g of NKA beads was prewetted with 40 mL of ethanol. Then immobilization of lipase AYS on the NKA 5143

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from GL Science Inc. (Tokyo, Japan) was adopted to separate and detect the targeted products. Injection volume was 10 μL. The mobile phase was acetonitrile/2-propanol (65:35, v/v). Column temperature was 40 °C, and flow rate was 1.0 mL/min. Detection was carried out by PDA at 205 nm. A 50 μL aliquot of the reaction product was taken from the reaction system and isolated by thin layer chromatography (TLC) plates with hexane/acetic ether/acetic acid (88:12:1, v/v/v) as the developing solvent.24,25 First, products were diluted with the developing solvent and spotted on TLC plates repeatedly (>10 times), with standard substances of phytosterol ester and diglyceride as references. Then development was carried out in the developing solvent, and the spots were visualized by iodine vapor and exposure in an oven set at 60 °C for 5 min. Ten bands corresponding to diglycerides and phytosterol esters were scraped off and collected, respectively. Fatty Acid Composition Analysis. The fatty acid compositions of the triglycerides, diglycerides, and phytosterol esters were further confirmed by gas chromatography (GC) analysis. First, the fatty acid methyl esters (FAMEs) were prepared from diglycerides, triglycerides, and phytosterol esters according to “Animal and Vegetable Fats and Oils Preparation of FAMEs” (GB/T 17376, ISO 5509:2000, IDT) with some modification. Briefly, 100 μL of sodium methylate solution (0.5 M in methanol) and 2.5 mL of hexane were added to 0.2 g of triglyceride or a certain amount of other targets (diglycerides or phytosterol esters; each needs 10 bands collected from TLC) and then reacted for 10 min in the shaker at 55 °C. The FAMEs were extracted twice with 20 mL of hexane, and after 10 min of centrifugation, the supernatant FAMEs were concentrated with nitrogen before being preserved for GC analysis. The fatty acid compositions of the diglycerides, triglycerides, and phytosterol esters in the products were analyzed by GC. An Agilent 7890 series gas chromatograph (Hewlett-Packard Co., Avondale, PA, USA), equipped with a flame ionization detector (FID) and a fused silica capillary column (DB-5 HT, 30.0 m × 320 μm × 0.25 μm, Agilent Technologies, Palo Alto, CA, USA) was used. The injection volume was 1 μL at a split ratio of 80:1. The carrier gas was nitrogen, and the nitrogen flow rate was 1.5 mL/min in constant flow rate mode. The flow rates of hydrogen and air were 40 and 400 mL/min, respectively, and the makeup gas was also nitrogen at a flow rate of 41.5 mL/min. The injector and detector temperatures were maintained at 250 and 300 °C, respectively. The oven temperature was held at 210 °C for 9.0 min, then increased to 250 °C at a rate of 20 °C/min, and held at 250 °C for 5 min. The fatty acid compositions of the diglycerides, triglycerides, and phytosterol esters in the products were compared with the standard substances of different fatty acids and confirmed by the retention time. The proportion of each fatty acid could be calculated by the ratio of peak area assigned to corresponding fatty acid in the GC profile. Physiochemical Properties of Functional Oils from Different Triglyceride Sources. The peroxide value (PV) and acid value (AV) were determined according to the National Standard of the People’s Republic of China (PRC) (GB/T 5538, ISO 3960:2001, IDT; GB/T 5530, ISO 660:1996, IDT).26 The conjugated diene (CD) value was determined as reported previously.27 Briefly, the sample was dissolved in 50 mL of cyclohexane and the optical density (1 cm light path) recorded at 234 nm against a cyclohexane blank. The conjugated diene value was calculated according to the equation

CD (mmol/kg of oil) =

AV × 100 εbm

biocatalyst for transesterification of phytosterol with triglyceride. Lipase AYS was the most effective among the enzymes tested (conversion >52%). Thus, lipase AYS was selected for the following experiments. To take advantage of high specific surface area and ideal porous structure, NKA beads were used as a support matrix to immobilize lipase AYS. However, it is difficult for the enzyme solution to access the inner surface of the NKA bead pores because of the highly hydrophobic nature of these surfaces. When prewetted with ethanol, the NKA beads were perfectly suspended in the enzyme solution, which could help the lipase reach the internal of the hydrophobic pores. The presence of ethanol may also contribute to a rearrangement of the secondary structure involving a higher accessibility of some hydrophobic side chains, which could increase the activity of noncovalently immobilized lipase.28 Furthermore, the catalytic activity of soluble lipase in the presence of 10% ethanol concentrations could not have been negatively affected.28 Thus, ethanol (10%, v/v) was added and acted as an intermediate between the lipase solution and the NKA beads in this experiment. The AYS@NKA was characterized by measuring the texture parameters of samples using a Brunauer−Emmett−Teller (BET) treatment of the N2 sorption isotherm data. It can be found that the surface area and pore volume of NKA are 562 m2/g and 0.66 cm3/g, respectively. After lipase absorption, the NKA beads have a lower surface area (501 m2/g) and a lower pore volume (0.63 cm3/g). The decrease of surface area and pore volume values of NKA after absorption indicated that the lipase AYS has been absorbed on the NKA surface and partly occupied the NKA surface pores. To evaluate the loading capacity of the carrier, 20 g of NKA beads was loaded with 400 mL of different initial lipase concentrations ranging from 0.5 to 7.0 mg/mL. The adsorbed AYS amount increased with increasing initial AYS amount until 6.5 mg/mL and reached a maximum value of 51.2 mg/g, which was higher than that of lipase immobilized on magnetic microspheres (48.1 mg/g) and a fibrous polymer (44.7 mg/ g).23,29 The result shows that the macroporous resin NKA beads have excellent adsorption properties and allow the adsorption of a high amount of AYS with high activity. Activity and Thermal Stability of Free AYS and AYS@ NKA. The catalytic activity and thermal stability of free AYS and AYS@NKA during the transesterification reaction were evaluated. As shown in Figure 2, almost no formation of phytosterol esters occurred in the absence of AYS. Under the temperature of 50 °C, the conversion of phytosterol esters increased rapidly during the first 12 h and reached a relatively high conversion (92.4%) with AYS@NKA, whereas the conversions of phytosterol esters reached only 68.6% after 24 h of reaction using free AYS as the catalyst. These facts indicated that the immobilized AYS@NKA showed much higher catalytic activity than that of free AYS, especially at the first 12 h. The high catalytic activity of the immobilized AYS@ NKA might be ascribed to the better dispersion effect as well as the higher mass transfer rate of the immobilized AYS@NKA in the reaction system compared with that of the free AYS, which might easily become aggregated in the reaction system. The conversion of phytosterol esters catalyzed by free AYS in 60 °C showed a sharp decrease compared with that in 50 °C, which might be because the higher temperature would inhibit the activity of lipase AYS. In contrast, AYS@NKA retained most of its initial activity, and the conversion was 87.2% under the same conditions, proving enhanced thermostability for this

(2)

where A is the absorbance at 234 nm of the test sample, V is the constant volume, ε is the molar absorption coefficient, 26000 L/(mol· cm), b is the thickness of the absorption cell, and m is the sample mass.

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RESULTS AND DISCUSSION Preparation and Characterization of the AYS@NKA. Five enzymes including lipase PS, lipase AYS, Novozym 435, Lipozyme TL IM, and Lipozyme RM IM were selected as the 5144

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Figure 3. Effect of phytosterol concentration on the conversion of phytosterol esters in lipase-catalyzed reaction. Reaction conditions: 1:2 molar ratio of phytosterols to sunflower oil, 25 mg/mL AYS@NKA in hexane, 180 rpm, and 50 °C.

Figure 2. Activity and thermal stability of AYS@NKA and free AYS. Each experiment from Figures 2−6 was duplicated three times. The standard deviations are shown as error bars in each figure.

composite material. This is attributed to the stabilizing effect of the AYS immobilized on the NKA surface, which prevents extensive structural changes typical of thermal denaturation. The ability to retain enzyme activity at high temperatures provides several processing advantages, such as improved substrate solubility. Optimization of Enzymatic Transesterification Conditions. Generally speaking, the kind of organic solvent is a key factor for enzymatic transesterification, because it affects not only the activity and stability of enzymes but also the solubility of substrates. The more hydrophobic solvent used, the greater activity and stability the lipase exhibited. Taking the whole into consideration, n-hexane, iso-octane, cyclohexane, and tert-amyl alcohol, which show different log P values, were chosen as solvents for the esterification reaction. As shown in Table 1, the

extent.29 With phytosterol concentrations of 50 and 80 mmol/ L, the phytosterol esters reached relatively high conversions, 99.8 and 91.5% after 12 h, respectively. Considering both the conversion and the economical aspect of the process, a relatively higher phytosterol concentration of 80 mmol/L was selected for the subsequent experiment. The effect of the molar ratio of phytosterols to triglycerides on the conversion of phytosterol esters was evaluated. The molar ratio of phytosterols to sunflower oil was 1:1−1:8. With the molar ratio of phytosterols to sunflower oil increasing from 1:1 to 1:8, as shown in Figure 4, the conversions increased from

Table 1. Effects of Organic Solvents on Conversion of Enzymatic Transesterificationa solvent

log P

conversion (%)

n-hexane iso-octane cyclohexane tert-amyl alcohol

3.2 4.5 3.5 1.2

95.2 89.7 75.4 80.6

a

Reaction conditions: 80 mmol/mL phytosterols, 1:2 molar ratio of phytosterols to sunflower oil, 25 mg/mL AYS@NKA in solvent, 180 rpm, and 50 °C.

reaction in hexane exhibits the highest conversion after 12 h among the four solvents. As is known to all, the conversion is affected by both the solubility of substrates and solvent polarity. The highest conversion in hexane might ascribed to the compromise of the solubility of substrates and solvent polarity. The effect of the substrate concentration on the conversion of phytosterol esters was evaluated. The molar ratio of phytosterols to sunflower oil was 1:2. As shown in Figure 3, it was observed that increasing the concentration of phytosterols from 50 to 150 mmol/L led to a continued decrease in the conversion. Some of the phytosterols were not soluble in the solvent at higher concentration, which was responsible for the lower conversion. In addition, an increase in phytosterols and triglyceride concentration may change the catalytic environment and the active site of AYS@NKA to some

Figure 4. Effect of molar ratio of phytosterols to triglycerides on the conversion of phytosterol esters in lipase-catalyzed reaction. The phytosterol concentration was 80 mmol/L. Other conditions were the same as described in Figure 3.

91.8 to 97.4% and then decreased to 84.3%. This might due to the fact that an increase in triglyceride molar ratio may not only change the catalytic environment and the active site of AYS@ NKA but also increase the viscosity of the reaction solvent and thus decrease the mass transfer rate.30 Considering the extent of esterification and economical aspect of the process, the mole ratio of 1:2 was selected for the subsequent experiment. The influence of the AYS@NKA load on the conversion of phytosterol esters was evaluated with a 1:2 phytosterols to sunflower oil molar ratio and an 80 mmol/L phytosterol 5145

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concentration using various amounts of AYS@NKA from 0 to 40 mg/mL. The content of phytosterol esters was very low (18.7%) in the absence of AYS@NKA (See Figure 2). From Figure 5, it observed that with the amount of AYS@NKA

Figure 6. Effect of reaction temperature on the conversion of phytosterol esters in lipase-catalyzed reaction. Reaction conditions: 80 mmol/L phytosterols, 1:4 molar ratio of phytosterols to sunflower oil, hexane, and 180 rpm. Figure 5. Effect of AYS@NKA load on the conversion of phytosterol esters in lipase-catalyzed reaction. Reaction conditions: 80 mmol/L phytosterols, 1:4 of the molar ratio of phytosterols to sunflower oil, hexane, 180 rpm, and 50 °C.

reaction medium.31 HPLC analysis results showed that the peaks of two functional ingredients including phytosterol esters and diglycerides were observed in two functional oils obviously (Figuer 7). The peaks of phytosterol in functional oils were almost disappeared, which prove that the enzymatic transesterification ratio was quite high (>92.1%). As listed in Table 2, the contents of functional ingredients in four different functional oils were in the ranges of 10.4−12.5% for diglycerides and 20.2−25.1% for phytosterol esters, respectively. Table 2 also lists the FA profile of the original vegetable oils (four kinds of triglycerides) and phytosterol esters and diglycerides produced. Four vegetable oils, which cover a wide range of oleic, linoleic, and linolenic acid values, were used as the sources of different acyl donors. Analysis of the FA composition of vegetable oils, phytosterol esters, and diglycerides by GC showed that almost all of the FAs present in the vegetable oils were incorporated into the corresponding phytosterol esters and diglycerides. In other words, the developed method could produce functional oils rich in both phytosterol esters and diglycerides with controllable FA composition by modulating the initial acyl donors with the wanted FA composition. Physiochemical Properties of Functional Oils Prepared from Different Vegetable Oils. The physiochemical properties of functional oils prepared from different vegetable oils are shown in Table 3. The PV and CD values were low, which showed that few products of peroxidation were produced, a benefit of the mild reaction conditions. The AV and PV values were less than 1.0 mg KOH/g and 2.1 mmol/kg, respectively, which agree with the standards of sunflower oil, rapeseed oil, corn oil, and linseed oil of China (GB10464-2003, GB1536-2004, GB19111-2003, GB8235-2008) as well as the cold pressed oils standards of the Codex Alimentarius Commission (CAC) and thus could be used as the functional lipid alone or as the fat-soluble additive in vegetable oils. Conclusion. In conclusion, a rapid and convenient esterification method using immobilized AYS@NKA as catalyst was developed to synthesize novel functional oils rich in both phytosterol esters and diglycerides in high yield under mild conditions. Furthermore, the developed method could produce phytosterol esters and diglycerides with controllable FA

loaded increasing from 10 mg/mL to 40 mg/mL, the conversion increased from 78.1 to 96.1%. A good synthesis method should consider the conversion and economical interest of the reaction, in other words, use less AYS@NKA to obtain a satisfactory transesterification ratio. Using a minimal amount of AYS@NKA such as 10 mg/mL would be economically attractive, but conversion of phytosterols reached only 78.1% after 12 h. Increasing the amount of AYS@NKA loaded led to better production of phytosterol esters. The formation of the phytosterol esters was much higher with a 25 mg/mL AYS@NKA load and reached a 93.2% conversion after 12 h of reaction. Considering both the conversion and the economical aspects of the process, 25 mg/mL AYS@NKA was selected for the subsequent experiment. The effect of temperature on the conversion of phytosterol esters was evaluated. The temperature ranged from 40 to 60 °C. As the reaction temperature increased from 40 to 60 °C, as shown in Figure 6, the conversion of phytosterol esters increased from 78.7 to 97.4% and then decreased to 79.2%. The highest conversion was achieved when the temperature was 50 °C. As the temperature was up to 60 °C, the conversion showed an obvious decrease (79.2%), and the energy consumption was high as well. Thus, 50 °C was selected as the reaction temperature for the subsequent experiment, which yielded an excellent conversion as well as energy savings. Synthesis of Functional Oils with Different Vegetable Oils. Under the optimized conditions, the transesterification of phytosterols with four vegetable oils including sunflower oil, corn oil, rapeseed oil, and linseed oil was investigated. As listed in Table 2, the transesterification ratios of phytosterols with four vegetables oils were between 92.1 and 100.2%. The results suggest that AYS@NKA could be used to catalyze the transesterification of phytosterols with different acyl donors with relatively high conversion. This might be ascribed to the fact that lipases prefer to adsorb on hydrophobic supports, involving the adsorption of the hydrophobic areas surrounding the active center and leaving the active site fully exposed to the 5146

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Table 2. Fatty Acid Profile of the Phytosterol Esters (PE) and Diglycerides (DG) in Functional Oils Corresponding to Different Vegetable Oils as the Acyl Donorsa fatty acid composition (%, w/w) product sunflower oil functional oil 1

corn oil functional oil 2

rapeseed oil functional oil 3

linseed oil functional oil 4

content (%)

C16:0

C18:0

C18:1

C18:2

C18:3

conversion (%)

PE DG

25.1 12.3

6.4 6.9 5.3

3.4 2.4 5.4

30.4 29.4 32.4

59.8 61.3 56.9

0 0 0

98.9

PE DG

20.2 10.4

13.0 13.5 12.0

1.7 2.6 0

31.1 24.8 42.7

54.2 59.1 45.3

0 0 0

96.5

PE DG

24.1 12.5

4.9 4.2 6.9

1.9 2.6 0

64.9 64.8 65.2

20.9 20.3 22.5

7.4 8.1 5.5

92.1

PE DG

22.3 11.2

5.2 7.4 0

3.2 4.5 0

18.1 14.5 26.8

16.3 15.3 18.7

57.2 58.3 54.6

100.2

a Values are the average of analysis from triplicate sets. Functional oils 1−4 were obtained by transesterification of phytosterols with triglycerides from sunflower oil, corn oil, rapeseed oil, and linseed oil, respectively. Contents of PE and DG were short for the content of phytosterol esters and diglycerides in different kinds of functional oils. Reaction conditions: 80 mmol/L phytosterols, 1:2 molar ratio of phytosterols to triglycerides, nhexane, 180 rpm, and 50 °C.

Figure 7. Liquid chromatograms of sunflower oil (A1), corn oil (B3), and responding functional oils (A2, B4). Peaks: (1) diglycerides; (2) phytosterols; (3) triglycerides; (4) phytosterol esters.

FA composition of the corresponding triglycerides. These findings could promote the wide application of the novel functional oils produced by the food grade process in different formulations of functional foods.

Table 3. Physiochemical Properties of Four Vegetables and Corresponding Functional Oils product sunflower oil functional oil corn oil functional oil rapeseed oil functional oil linseed oil functional oil

1 2 3 4

PVa (mmol/kg)

AVa (mg of KOH/g)

CDa (mmol/kg)

1.6 1.6 1.5 1.8 1.5 1.7 1.8 2.1

0.8 0.9 0.5 0.7 0.6 0.8 0.7 1.0

1.93 1.87 1.47 1.62 1.96 1.95 1.72 1.69

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AUTHOR INFORMATION

Corresponding Author

*(F.-H.H.) Phone: +86-27-86827874. Fax: +86-27-86815916. E-mail: [email protected]. Author Contributions ∥

a

M.-M.Z. and Q.H. contributed equally to this work.

Funding

PV, AV, and CD are abbreviations for peroxide value, acid value, and conjugated diene value, respectively. Functional oils 1−4 are the same as those described in Table 2.

This work was supported by the National Science Foundation of China (NSFC-31371843, 31101353), the Director Fund of Oil Crops Research Institute (1610172014006), the Open Foundation of Hubei Key Laboratory of Lipid Chemistry and Nutrition (2012003), and the Foundation for Modern Agroindustry Technology Research System (CARS-13).

composition by modulating the initial acyl donors with the wanted FA composition. The physiochemical properties of four prepared functional oils were found to be closely related to the 5147

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Notes

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The authors declare no competing financial interest.

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dx.doi.org/10.1021/jf500744n | J. Agric. Food Chem. 2014, 62, 5142−5148