Article pubs.acs.org/JAFC
Impact of Association Colloids on Lipid Oxidation in Triacylglycerols and Fatty Acid Ethyl Esters Rika Homma,*,†,‡ Karin Suzuki,† Leqi Cui,† David Julian McClements,†,§ and Eric A. Decker†,§ †
Department of Food Science, University of Massachusetts Amherst, 102 Holdsworth Way, 230 Chenoweth Laboratory, Amherst, Massachusetts 01003, United States ‡ Health Care Food, Kao Corporation, 2-1-3 Bunka, Sumida-ku, Tokyo 1318501, Japan § Bioactive Natural Products Research Group, Department of Biochemistry, Faculty of Science, King Abdulaziz University, P.O. Box 80203, Jeddah 21589, Saudi Arabia ABSTRACT: The impact of association colloids on lipid oxidation in triacylglycerols and fatty acid ethyl esters was investigated. Association colloids did not affect lipid oxidation of high oleic safflower and high linoleic safflower triacylglycerols, but were prooxidative in fish triacylglycerols. Association colloids retarded aldehyde formation in stripped ethyl oleate, linoleate, and fish oil ethyl esters. Interfacial tension revealed that lipid hydroperoxides were surface active in the presence of the surfactants found in association colloids. The lipid hydroperoxides from ethyl esters were less surface active than triacylglycerol hydroperoxides. Stripping decreased iron and copper concentrations in all oils, but more so in fatty acid ethyl esters. The combination of lower hydroperoxide surface activity and low metal concentrations could explain why association colloids inhibited lipid oxidation in fatty acid ethyl esters. This research suggests that association colloids could be used as an antioxidant technology in fatty acid ethyl esters. KEYWORDS: association colloids, critical micelle concentration, lipid oxidation, lipid hydroperoxides, interfacial tension, prooxidant transition metals
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INTRODUCTION Lipid oxidation is a major cause of food quality deterioration, and has been a challenge for manufacturers and food scientists. Lipid oxidation leads not only to nutrient loss but also causes color alterations and off-flavor generation, and thus decreases product quality. To help reduce the risk of heart disease, it has been suggested to replace solid fats with oils rich in polyunsaturated fatty acids. In general, oxidation rates increase with the increasing oil unsaturation. When foods are formulated with high levels of polyunsaturated fatty acids to improve the nutritional quality, control of lipid oxidation becomes a major challenge for the food industry. Many researchers have identified lipid oxidation mechanisms in foods.1−3 However, the mechanism of lipid oxidation in oil is still not completely understood, especially the physical and the chemical effects of surface-active minor components, which always exist even in refined oils. Oils contain not only triacylglycerols, but also various minor compounds such as diacylglycerols, monoacylglycerols, free fatty acids, tocopherols, sterols, phospholipids, and even water.4 Surface-active minor components have been reported to lower interfacial tension in oils, suggesting that they are concentrated at the oil−water interface and form association colloids. When surface-active components form association colloids, they can change the physical and the chemical properties of oil, which could impact lipid oxidation.5−8 Previously, our group has demonstrated the impact of association colloids on lipid oxidation. Diolelylphosphatidylcholine (DOPC), an example of a surface-active minor component in natural oils, formed association colloids and promoted the formation of both primary and secondary lipid © XXXX American Chemical Society
oxidations in stripped soybean oil, stripped soybean oil/ medium chain triacylglycerol (MCT) mixtures, and stripped corn oil/MCT mixtures.9−13 Mixtures of surface-active components (DOPC, dioleoylphosphatidylethanolamine, diolein, oleic acid, and stigmasterol) also formed association colloids in stripped corn oil/MCT mixture and promoted lipid oxidation when the concentrations of the mixed surface-active components were above the critical micelle concentration (CMC).14 Chen et al. proposed that association colloids reduced the oxidative stability of oil by attracting lipid hydroperoxides and metals ions into the oil−water interface.11 However, research has mainly focused on oils that mainly contain linoleic acid and have not studied oils where monounsaturated and omega-3 fatty acids predominate. Thus, to more fully understand the role of association colloids on the oxidation of triacylglycerols, future studies using different fatty acid compositions are needed. Lipid oxidation is an issue for not only triacylglycerols, but also all other fatty acid related compounds such as fatty acid ethyl esters. Recently, fish oil ethyl ester has become a popular dietary supplement due to their high levels of docosahexaenoic acid (DHA) and eicosapentaenoic acid (EPA).15,16 There has been a lot of research conducted on lipid oxidation rates of fatty acid ethyl esters in bulk oil and emulsions.17−20 Unfortunately, the impact of association colloids on oxidative stability of fatty acid ethyl esters is largely unknown. Received: August 3, 2015 Revised: October 16, 2015 Accepted: October 27, 2015
A
DOI: 10.1021/acs.jafc.5b03807 J. Agric. Food Chem. XXXX, XXX, XXX−XXX
Article
Journal of Agricultural and Food Chemistry Here, the first purpose of this study was to investigate the effects of the fatty acids composition on the formation of association colloids in triacylglycerols oils and fatty acid ethyl esters and to see how the association colloids impacted lipid oxidation kinetics. Further, we investigated the ability of different fatty acid lipid hydroperoxides to decrease interfacial tension and how stripping oils of their minor components impacted metal concentrations and lipid oxidation to better understand lipid oxidation mechanism. This research provides a better understanding of how association colloids can impact the lipid oxidation of oils, which in turn could provide important information on how to control oxidative rancidity.
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by GC without any preparation. Table 1 shows major fatty acids content in the oils that we used in this study.
Table 1. Major Fatty Acid Content in Oila oil
OA
LA
ALA
EPA
DHA
HOSO HLSO fish oil ethyl oleate ethyl linoleate fish oil ethyl ester
77.7% 16.6% 10.6% 99.1% 0.1% 9.2%
13.6% 71.0% 1.2% 0.1% 99.3% 0.9%
n.d. n.d. 0.9% 0.2% 0.3% 1.0%
n.d. n.d. 39.7% n.d. n.d. 19.8%
n.d. n.d. 22.5% n.d. n.d. 13.3%
a Oleic acid (OA), linoleic acid (LA), α-linolenic acid (ALA), eicosapentaenoic acid (EPA), docosahexaenoic acid (DHA).
MATERIALS AND METHODS
Materials. High oleic safflower oil (HOSO) and high linoleic safflower oil (HLSO) were purchased from the Nissin Oillio Group, Ltd. (Chuo-ku, Tokyo, Japan). Ethyl oleate and ethyl linoleate were purchased from NU-CHECK-PREP, Inc. (Elysian, MN). Fish oil ethyl ester and fish oil were provided by BASF SE (Ludwigshafen, Germany). 1,2-Dioleoyl-sn-glycero-3-phosphatidylcholine (DOPC), 1,2-dioleoyl-sn-glycero-3-phosphatidylethanolamine (DOPE), 1,2-dioleoyl-sn-glycerol (DAG), β-sitosterol, oleic acid, and 7,7,8,8tetracyanoquinodimethane (TCNQ) were purchased from SigmaAldrich Co. (St. Louis, MO). Rose Bengal and hexane were purchased from Fisher Scientific (Fair Lawn, NJ). Medium chained triacylglycerols (MCT) were obtained from Sasol North America Inc. (Huston, TX). All chemicals and solvents were used without further purification. Deionized water was used in all experiments. Glassware was submerged in 2 M HCl overnight to remove metals, followed by rinsing with deionized water. Preparation of Stripped Oil. To avoid the influence of the minor compounds such as tocopherols, phospholipids, free fatty acids, and mono- and diacylglycerols normally present in commercial oil, stripped oils were prepared as described by Kittipongpittaya et al.12 Briefly, a chromatographic column (3.0 cm diameter, 35 cm long) was used to isolate fatty acid ethyl esters or triacylglycerols. Three layers were packed into the column sequentially. The bottom layer was packed with 22.5 g of silicic acid (washed with distilled and deionized water and activated at 110 °C for 48 h). Activated charcoal (5.6 g) was then used for the middle layer and another 22.5 g of silicic acid for the top layer. The different oil types (30 g) were mixed with 30 mL of hexane, and the mixture was passed through the column using 270 mL of hexane for elution. The solvent was then removed by a vacuum rotary evaporator (rotary evaporator R 110, Buchi, Flawil, Switzerland) at 25 °C, and the remaining solvent was evaporated by nitrogen flushing. HPLC analysis detected no tocopherols in stripped oils (method described below). The water content of all triacylglycerols and fatty acid ethyl esters was less than 100 ppm as determined by Karl Fischer analysis using an 831 KF Coulometer (Metrohm, FL). Tocopherols Concentration Measurement. Oil samples (50 mg) were dissolved in 1 mL of isopropanol and 1 mL of methanol and then passed through a 0.2 μm filter (Fisher Scientific, PA). Sample (20 μL) was then injected into a Shimadzu HPLC system equipped with a Beckman Ultrasphere C18 reverse phase column (150 mm × 4.6 mm, 5 μm). The mobile phase was isocratic methanol at a flow rate of 1 mL/min. A Waters 474 scanning fluorescent detector (Waters, MA) was used to detect tocopherols at an excitation wavelength of 290 nm and an emission wavelength of 330 nm. Peak integration was performed using Shimadzu EZstart software (version 7.2). Tocopherols in the samples were identified and quantified by comparing their relative retention times and peak areas with authentic compounds. Determination of the Fatty Acid Composition of Triacylglycerols. Fatty acid methyl esters were prepared in accordance with “Methods of preparing fatty acid methyl esters (2.4.1 − 1996)” in “Standard methods for the Analysis of Fats, Oils and Related Materials” edited by Japan Oil Chemists Society. The resulting samples were measured by the American Oil Chemists Society official method Ce 1f-96 (GC method). Fatty acid ethyl esters were measured
Determination of the Critical Micelle Concentration. The critical micelle concentrations (CMC) of stripped oils with mixed surface-active components were determined by using the TCNQ solubilization technique.12,21 Briefly, the absorbance of TCNQ in the oil (including the surface-active components DOPC, DOPE, DAG, βsitosterol, and oleic acid mixture at a molar ratio of 3.78:0.67:2.25:0.97:0.43) was determined at concentrations ranging from 1 to 1000 μmol/kg oil. The ratios of the surface-active components are similar to those found in refined corn oil.4,22 The oils containing surface-active components were stirred at 400 rpm at 20, 37, and 55 °C for 12 h prior to adding 1 mg of TCNQ/g oil and mixing for another 3 h. The excess TCNQ was removed by centrifugation at 3000g for 20 min. The absorbance was measured at 480 nm using a spectrophotometer (Shimadzu 2014, Tokyo, Japan). The CMC was determined as the inflection point in the semilog plot of absorbance versus surface-active components concentration as described previously.12 Sample Preparation for Oxidation Studies. A mixture of surface-active components including DOPC, DOPE, DAG, βsitosterol, and oleic acid at a molar ratio of 3.78:0.67:2.25:0.97:0.43 was added to the stripped oil at concentrations below (10 μmol/kg oil) or above (500 or 1000 μmol/kg oil) the CMC of the different oil types. The samples were then stirred at 20 °C for 12 h. For the lipid oxidation studies, 1.0 mL samples were aliquoted into 10 mL headspace vials, sealed with aluminum caps with PTFE/silicone septa, and stored in dark at 20 °C (ethyl linoleate, fish oil ethyl ester, and fish oil), 37 °C (ethyl oleate and high linoleic safflower oil), and 55 °C (high oleic safflower oil). These storage temperatures were selected by preliminary oxidation studies, during which each stripped oil showed an oxidation lag phase of more than 2 days allowing for comparison of the effect of association colloids on oxidation lag phase. Lag phase is an important parameter of the shelf life of oils because rancidity can be detected after the lag phase. Measurement of Lipid Oxidation. Lipid oxidation was determined by monitoring lipid hydroperoxides and headspace aldehyde (propanal, hexanal, and nonanal) formation. Lipid hydroperoxides were measured as primary oxidation products according to Kittipongpittaya et al.12 Briefly, 2.8 mL of a mixture of methanol/ butanol (2:1, v/v) was added to oil samples of known weight (15−25 mg). Next, 15 μL of 3.94 M ammonium thiocyanate and 15 μL of 0.072 M Fe2+ (ferrous sulfate) were added, and the solution was vortexed. After a 20 min reaction time, the absorbance was measured at 510 nm using a Genesys 20 spectrophotometer (Thermo Spectronic, Waltham, MA). The concentration of lipid hydroperoxides was calculated from a standard curve prepared from cumene hydroperoxide ranging from 0.7 to 17 mM. Samples with lipid hydroperoxides concentrations higher than 17 mM were diluted with methanol/butanol (2:1, v/v) before measurements. The secondary lipid oxidation products propanal, hexanal, and nonanal were measured as oxidation products of omega-3, -6, and -9 fatty acids, respectively. Aldehydes were quantified by gas chromatography (GC, model GC-2014, Shimadzu, Tokyo, Japan) equipped with an auto sampler and flame ionization detector (FID) as in Chen et al.9 B
DOI: 10.1021/acs.jafc.5b03807 J. Agric. Food Chem. XXXX, XXX, XXX−XXX
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Samples (1 mL) were incubated at 55 °C for 10 min to allow volatile compounds to enter the headspace of the 10 mL headspace gas chromatography vial. A divinylbenzene/carboxen/polydimethylsiloxane (DVB/carboxen/PDMS) solid-phase micro extraction (SPME) fiber (50/30 μm, Supelco, Bellefonte, PA) was exposed to the headspace above the sample for 2 min to adsorb volatile components, followed by desorption for 3 min at 250 °C in the GC injector at a split ratio of 1:7. Volatile compounds were separated on a fused-silica capillary column (30 m × 0.32 mm inner diameter × 1 μm) coated with 100% polydimethylsiloxane (Equity-1, Supelco). For propanal and hexanal analysis, the run time was 10 min per sample with temperatures of 250, 65, and 250 °C for the injector, oven, and detector, respectively. For nonanal analysis, the run time was 25 min per sample with temperatures of 250, 90, and 250 °C for the injector, oven, and detector, respectively. Propanal, hexanal, and nonanal concentrations were calculated from the peak areas using a standard curve made from MCT containing known propanal, hexanal, and nonanal concentrations from 2 to 1000 μM. The lag phase is defined as the length of time before rapid acceleration of lipid oxidation. Lag phase was determined by drawing tangents to the lines of the lag and exponential phases in the plot of the lipid hydroperoxides concentrations or aldehyde concentrations versus storage days.23 Formation of Lipid Hydroperoxides. Lipids hydroperoxides were produced from stripped high oleic safflower oil, high linoleic safflower oil, fish oil, ethyl oleate, ethyl linoleate, and fish oil ethyl ester by mixing 1 g of stripped oil and 0.02 g of Rose Bengal, a singlet oxygen generator.24 The mixture was incubated at 5 °C and illuminated with a 60 W (9000 Lux) lamp for different periods of time (high oleic safflower oil, 41 h; high linoleic safflower oil, 28 h; fish oil, 24 h; ethyl oleate, 44 h; ethyl linoleate, 20 h; fish oil ethyl ester, 20 h) to produce lipid hydroperoxides with concentration of 100−300 mmol/kg oil (high oleic safflower oil, 298 mmol/kg oil; high linoleic safflower oil, 297 mmol/kg oil; fish oil, 194 mmol/kg oil; ethyl oleate, 273 mmol/kg oil; ethyl linoleate, 294 mmol/kg oil; fish oil ethyl ester, 174 mmol/kg oil) without the generation of aldehydes. For interfacial tension measurement, lipid hydroperoxides were diluted to 50 mmol/ kg oil in freshly stripped oils. Headspace aldehydes were not detected in any samples, indicating that minimal lipid hydroperoxides breakdown occurred during lipid hydroperoxide formation. Interfacial Tension Measurements. The interfacial tension at the oil−water interface was measured using a droplet shape analysis device (DSA 100, Krüss GmbH, Hamburg, Germany). A straight needle with diameter of 1.07 mm was used to create a water drop in the oil in a quartz cell. Each sample was a composite of measurements made every 0.2 s for 5 min. Interfacial tension values were calculated on the basis of the Young−Laplace equation by a drop shape analysis program supplied by the instrument manufacture. Metal Concentration Measurements. To quantify iron concentrations, oils were subjected to ICP-MS analysis. Oils were prepared for ICP-MS using a method as described in Savio et al.25 Approximately 500 mg of oil was weighed directly into microwave digestion vessels, followed by addition of nitric acid (5.0 mL). Microwave digestion was carried out by a Mars Xpress (CEM, Matthews, NC) by bringing the temperature to 180 °C in 10 min and then holding at 180 °C for an additional 20 min. Digested samples were then diluted with deionized water and held at 4 °C until analysis. ICP-MS analyses were performed on a PerkinElmer NexION 300X ICP mass spectrometer, and 197Au was measured under standard mode.26 Operating conditions are listed as below: nebulizer flow rate, 0.95−1 L/min; rf power, 1600 W; plasma Ar flow rate, 18 L/min; dwell time, 50 ms. A series of standard solutions (gold concentration: 20, 10, 5, 2, 1, 0.5, 0.2, 0 ppb) was prepared for each experiment. Statistical Analysis. All experiments used triplicate samples (three samples taken at each time point), and each experiment was repeated at least two times. Data were presented as mean values ± standard deviation. Data results were analyzed by analysis of variance (ANOVA) using SPSS 14.0 (SPSS Inc., Chicago, IL).
RESULTS AND DISCUSSION Critical Micelle Concentration of Mixed SurfaceActive Components in Stripped Oils. In the presence of Table 2. Water Content (ppm) and Critical Micelle Concentrations (CMC, μmol/kg oil) of Mixed SurfaceActive Components (DOPC, DOPE, DAG, β-Sitosterol, and Oleic Acid Mixture at a Molar Ratio of 3.78:0.67:2.25:0.97:0.43, Respectively) in Stripped Oil CMC (μmol/kg oil) stripped oil
water content (ppm) 20 °C
20 °C
37 °C
55 °C
HOSO HLSO fish oil ethyl oleate ethyl linoleate fish oil ethyl ester
59 39 77 59 84 61
55 100 550 32 80 165
28 69 267 17 55 84
12 35 79 no data no data no data
Figure 1. Formation of lipid hydroperoxides (a) and headspace nonanal (b) in stripped HOSO with mixed surface-active components (DOPC, DOPE, DAG, sitosterol, and oleic acid mixture at a molar ratio of 3.78:0.67:2.25:0.97:0.43, respectively) at 0, 10, and 500 μmol/ kg oil during storage at 55 °C. Values represent means ± standard deviation (n = 3).
a small amount of water naturally occurring in oils, surfaceactive components tend to form association colloids with their polar heads oriented toward the water core and their hydrophobic tails partitioning into the oil. The concentration C
DOI: 10.1021/acs.jafc.5b03807 J. Agric. Food Chem. XXXX, XXX, XXX−XXX
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Figure 2. Formation of lipid hydroperoxides (a) and headspace hexanal (b) in stripped HLSO with mixed surface-active components (DOPC, DOPE, DAG, sitosterol, and oleic acid mixture at a molar ratio of 3.78:0.67:2.25:0.97:0.43, respectively) at 0, 10, and 500 μmol/ kg oil during storage at 37 °C. Values represent means ± standard deviation (n = 3).
Figure 3. Formation of lipid hydroperoxides (a) and headspace propanal (b) in stripped fish oil with mixed surface-active components (DOPC, DOPE, DAG, sitosterol, and oleic acid mixture at a molar ratio of 3.78:0.67:2.25:0.97:0.43, respectively) at 0, 10, and 1000 μmol/kg oil during storage at 20 °C. Values represent means ± standard deviation (n = 3).
at which surface-active components begin to form association colloids is defined as a critical micelle concentration (CMC). The CMC of the mixture of the surface-active components, DOPC, DOPE, DAG, β-sitosterol, and oleic acid at a molar ratio of 3.78:0.67:2.25:0.97:0.43, respectively, was determined at 20, 37, and 55 °C. As shown in Table 2, mixed components were able to form association colloids in all oils used in this study. The CMC of mixed minor components in stripped high oleic safflower oil was 55 μmol/kg oil at 20 °C. As the degree of oil unsaturation increased from high oleic safflower oil to high linoleic safflower oil and fish oil, the CMC increased from 55 to 100 and 550 μmol/kg oil at 20 °C, respectively. The CMC values of the mixed minor components in stripped fatty acid ethyl esters were 32 μmol/kg oil (in stripped ethyl oleate), 80 μmol/kg oil (in stripped ethyl linoleate), and 165 μmol/kg oil (in stripped fish oil ethyl ester) at 20 °C. These results suggest that CMC increased with increasing degree of oil unsaturation. This could be due to the increasing polarity of the unsaturated fatty acids, which would decrease the driving forces for polar surface active compounds to orient away from the lipid phase by forming association colloids. Impact of Association Colloids on Oxidative Stability of Triacylglycerols. The concentrations of mixed surfaceactive components added to stripped high oleic safflower oil
and stripped high linoleic safflower oil were 10 and 500 μmol/ kg, and 10 and 1000 μmol/kg for stripped fish oil. These concentrations corresponded to levels below (10 μmol/kg oil) and above (500 and 1000 μmol/kg oil) the CMC in each stripped oil. Figures 1 and 2 show that the addition of 10 and 500 μmol/kg oil mixed surface-active components did not change the lag phases of both lipid hydroperoxides formation (primary oxidation) and nonanal or hexanal formation (secondary oxidation) in stripped high oleic safflower oil and high linoleic safflower oil. Stripped fish oil with 10 μmol/kg oil mixed surface-active components showed same lag phases of lipid hydroperoxides and propanal formation as compared to the control (Figure 3). However, once mixed surface-active components were added at 1000 μmol/kg in stripped fish oil, which was above the CMC, the lag phases of both lipid hydroperoxides and propanal formation decreased from 10 to 7 days. Our previous study showed that the mixture of surface-active components above the CMC accelerated the oxidation rate of a stripped corn oil/MCT (1/3) mixture.14 Reversed micelles formed by DOPC, DOPE, and their combination also accelerated the oxidation rate of soybean oil, a soybean oil/ MCT (1/3) mixture, and a corn oil/MCT (1/3) mixture.9−13 However, we also found that reversed micelle formed by D
DOI: 10.1021/acs.jafc.5b03807 J. Agric. Food Chem. XXXX, XXX, XXX−XXX
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Figure 4. Formation of lipid hydroperoxides (a) and headspace nonanal (b) in stripped ethyl oleate with mixed surface-active components (DOPC, DOPE, DAG, sitosterol, and oleic acid mixture at a molar ratio of 3.78:0.67:2.25:0.97:0.43, respectively) at 0, 10, and 500 μmol/kg oil during storage at 37 °C. Values represent means ± standard deviation (n = 3).
Figure 5. Formation of lipid hydroperoxides (a) and head space hexanal (b) in stripped ethyl linoleate with mixed surface-active components (DOPC, DOPE, DAG, sitosterol, and oleic acid mixture at a molar ratio of 3.78:0.67:2.25:0.97:0.43, respectively) at 0, 10, and 500 μmol/kg oil during storage at 20 °C. Values represent means ± standard deviation (n = 3).
DOPC did not accelerate the oxidation rate of corn oil.27 In this study, association colloids accelerated the oxidation rate of stripped fish oil, but did not affect the oxidation rate of stripped high oleic safflower oil and high linoleic safflower oil. These studies suggest that the impact of association colloids on oxidative stability of triacylglycerols differs depending on the oil. Impact of Association Colloids on Oxidative Stability of Fatty Acid Ethyl Esters. The concentrations of mixed surface-active components added to stripped fatty acid ethyl esters were 10 and 500 μmol/kg, which are above and below the CMC, respectively. The lag phases of both lipid hydroperoxides and nonanal formation in stripped ethyl oleate with 10 μmol/kg oil mixed surface-active components were similar to the control (Figure 4). However, when 500 μmol/kg oil mixed surface-active components was added to stripped ethyl oleate, the lag phase of lipid hydroperoxides formation increased from 13 to 25 days. Further, the lag phase of nonanal formation was increased from 11 to 21 days. The addition of 10 μmol/kg oil mixed surface-active components did not affect the lag phases of both lipid hydroperoxides and aldehyde formation in stripped ethyl linoleate and fish oil ethyl ester (Figures 5 and 6). On the other hand, 500 μmol/kg oil of mixed surface-active components increased the lag phase of hexanal formation in
stripped ethyl linoleate from 11 to 14 days, but did not affect the lag phase of lipid hydroperoxide formation. Stripped fish oil ethyl ester showed a trend similar to that of stripped ethyl linoleate. The addition of 500 μmol/kg oil mixed surface-active components increased the lag phase of propanal formation in stripped fish oil ethyl ester from 8 to 16 days, but did not affect the lag phase of lipid hydroperoxide formation. These results indicate that association colloids retarded the formation of secondary oxidation products in stripped fatty acid ethyl esters. The impact of mixtures of fatty acid ethyl ester was also investigated. Stripped fish oil ethyl ester (7 wt %) was added to stripped ethyl oleate, and the impact of association colloids on lipid oxidation was determined. The CMC of the mixed minor components in stripped ethyl oleate with 7 wt % fish oil ethyl ester was 50 μmol/kg oil. Surprisingly, the addition of 10 μmol/kg oil mixed surface-active components slightly decreased the lag phases of lipid hydroperoxides, propanal, and nonanal formation (Figure 7). When 500 μmol/kg oil mixed surface-active components were added, the lag phases of lipid hydroperoxides, propanal, and nonanal formation were significantly decreased from 15 to 7 days, 15 to 7 days, and 18 to 7 days, respectively. These results suggest that fish oil E
DOI: 10.1021/acs.jafc.5b03807 J. Agric. Food Chem. XXXX, XXX, XXX−XXX
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Figure 6. Formation of lipid hydroperoxides (a) and headspace propanal (b) in stripped fish oil ethyl ester with mixed surface-active components (DOPC, DOPE, DAG, sitosterol, and oleic acid mixture at a molar ratio of 3.78:0.67:2.25:0.97:0.43, respectively) at 0, 10, and 500 μmol/kg oil during storage at 20 °C. Values represent means ± standard deviation (n = 3).
ethyl ester could function as a prooxidant in ethyl oleate especially in the presence of association colloids. To better understand why association colloids impacted the oxidative stability of fatty acid ethyl esters differently from triacylglycerols, the surface activity of lipid hydroperoxides on different fatty acids and the levels of metal in oils after stripping were further investigated. Influence of Lipid Hydroperoxides on Interfacial Tension. The metal chelator, desferoxamine, can only inhibit lipid oxidation in bulk oil in the presence of association colloids, suggesting that association colloids increase the prooxidative activity of transition metals.11 This could occur because surface active compounds such as phospholipids can concentrate at the oil−water interface and produce a negative interfacial region that attracts metals.28 These interfacial transition metals could then interact with interfacial lipid hydroperoxides to increase lipid oxidation rates. Therefore, lipid hydroperoxides on different fatty acids could have different effects on lipid oxidation rates if the lipid hydroperoxides had different abilities to migrate to association colloids where they could be decomposed into free radicals. The ability of lipid hydroperoxides to alter oil−water interfacial tension was determined with triacylglycerols and fatty acid ethyl esters containing the mixed surface active components. As was expected, the mixed surface-active
Figure 7. Formation of lipid hydroperoxides (a), headspace propanal (b), and head space nonanal (c) in stripped ethyl oleate with 7% fish oil ethyl ester containing mixed surface-active components (DOPC, DOPE, DAG, sitosterol, and oleic acid mixture at a molar ratio of 3.78:0.67:2.25:0.97:0.43, respectively) at 0, 10, and 500 μmol/kg oil during storage at 20 °C. Values represent means ± standard deviation (n = 3).
components themselves reduced oil−water interfacial tension in the triacylglycerols (Table 3). All of the different types of triacylglycerols high in lipid hydroperoxides (e.g., stripped high oleic safflower oil, stripped high linoleic safflower oil, and stripped fish oil) also were able to reduce oil−water interfacial tension in the absence of the mixed surface active compounds. When triacylglycerols high in lipid hydroperoxides were added to the oils along with the mixed surface-active components, interfacial tension was reduced more than the individual F
DOI: 10.1021/acs.jafc.5b03807 J. Agric. Food Chem. XXXX, XXX, XXX−XXX
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Table 3. Interfacial Tension of Oil−Water Interface (mN/m) Containing Different Forms of Lipid Hydroperoxides and Mixed Surface-Active Components (DOPC, DOPE, DAG, Sitosterol, and Oleic Acid Mixture at a Molar Ratio of 3.78:0.67:2.25:0.97:0.43, Respectively) interfacial tension (mN/m) without mixed surface-active components
with mixed surface-active components
stripped oil
without hydroperoxide
with hydroperoxide
without hydroperoxide
with hydroperoxide
HOSO HLSO fish oil ethyl oleate ethyl linoleate fish oil ethyl ester ethyl oleate with 7% fish oil ethyl ester
27.9 32.7 31.7 30.4 30.3 32.8 30.4
26.9 28.3 26.6 30.0 29.4 30.0 25.0
25.1 29.1 28.4 30.3 29.9 32.5 30.3
21.6 20.5 12.8 27.5 24.0 21.4 20.5
interfacial tension reduction in stripped fish oil ethyl ester was larger than that in stripped ethyl linoleate and even more so than in stripped ethyl oleate, again suggesting that lipid hydroperoxide surface activity increased with increasing level of unsaturation of the fatty acids. However, the reductions of oil− water interfacial tension by stripped fatty acid ethyl esters high in lipid hydroperoxide were smaller than those by stripped triacylglycerols high in lipid hydroperoxides. This means that less fatty acid ethyl ester hydroperoxides would be at the oil− water interface of association colloids and thus might be less prooxidative because they would interact less with metals at the oil−water interface. Stripped ethyl oleate hydroperoxides had little impact on the interfacial tension of stripped ethyl oleate. On the other hand, the stripped fish oil ethyl ester hydroperoxides reduced interfacial tension when added to stripped ethyl oleate from 30.4 to 25.0 mN/m (without mixed surface-active components) and from 30.3 to 20.5 mN/m (with mixed surface-active components). These results suggest that the fish oil ethyl ester hydroperoxides have higher surface activity than the ethyl oleate hydroperoxides, and could partition at the oil−water interface in ethyl oleate solution. This could help explain why association colloids were not prooxidative in ethyl oleate or fish oil ethyl esters but were prooxidative in mixtures of fish oil ethyl ester and ethyl oleate. Effects of Transition Metals Concentration in Oils. The prooxidative activity of transition metals is strongly concentration dependent30 and thus could also help explain why we observed differences in lipid oxidation rates among the different lipid samples. This could be even more pronounced in stripped oils where the silicic acid and activated charcoal could remove metals. As expected, all oils used in this study contained iron and copper prior to stripping (Table 4). Stripping decreased iron and copper concentrations in all oils but more so in fatty acid ethyl esters where iron concentrations were reduced to below detectable levels. These very low transition metals concentrations in the fatty acid ethyl esters could help explain why the association colloids were not prooxidative as there would be low levels of interfacial iron. This would be especially true as fatty acid ethyl esters hydroperoxides were also less surface active than triacylglycerol hydroperoxides. The combination of low metals and lower lipid hydroperoxides at the interface of the association colloids would produce less free radicals from metal promoted lipid hydroperoxides decomposition. The research in this study presented a systematic investigation of three factors that could influence lipid oxidation in oils containing association colloids. These factors included
Table 4. Iron and Copper Concentrations (ppb) in Oil iron concentration (ppb) oil HOSO HLSO fish oil ethyl oleate ethyl linoleate fish oil ethyl ester
copper concentration (ppb)
before stripping
after stripping
before stripping
after stripping
1959 837 747 2013 315 458
587 667 554
