ProteinâProtein Multilayer Oil-in-Water Emulsions for the

Article pubs.acs.org/JAFC

Protein−Protein Multilayer Oil-in-Water Emulsions for the Microencapsulation of Flaxseed Oil: Effect of Whey and Fish Gelatin Concentration Patrick Fustier,* Allaoua Achouri, Ali R. Taherian, Michel Britten, Marylène Pelletier, Hassan Sabik, Sébastien Villeneuve, and Martin Mondor Food Research and Development Centre, Agriculture and Agri-Food Canada, 3600 Casavant Boulevard West, Saint-Hyacinthe, Quebec, Canada ABSTRACT: The impact of whey protein isolate (WPI) and fish gelatin (FG) deposited sequentially at concentrations of 0.1, 0.5, and 0.75% on the surface of primary oil-in-water emulsions containing 5% flaxseed oil stabilized with either 0.5% fish gelatin or whey protein, respectively, was investigated. The results revealed that the adsorption of WPI/FG or FG/WPI complexes to the emulsion interface led to the formation of oil-in-water (o/w) emulsions with different stabilities and different protection degrees of the flaxseed oil. Deposition of FG on the WPI primary emulsion increased the particle size (from 0.53 to 1.58 μm) and viscosity and decreased electronegativity (from −23.91 to −11.15 mV) of the complexes. Different trends were noted with the deposition of WPI on the FG primary emulsion, resulting in decreasing particle size and increasing electronegativity and viscosity to a lower extent. Due to the superior tension-active property of WPI, the amount of protein load in the WPI primary emulsion as well as in WPI/FG complex was significantly higher than the FG counterparts. A multilayer emulsion made with 0.5% WPI/ 0.75% FG exhibited the lowest oxidation among all of the multilayered emulsions tested (0.32 ppm of hexanal) after 21 days, likely due to the charge effect of FG that may prevent pro-oxidant metals to interact with the flaxseed oil. KEYWORDS: flax oil, multilayered oil-in-water emulsions, lipid oxidation, whey protein, gelatin

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INTRODUCTION Flaxseed oil represents a rich source of polyunsaturated fatty acids (PUFAs), which have been positively associated with a variety of human health benefits such as reducing the risk of coronary heart disease and diabetes, protection against inflammation, and prevention of certain types of cancer.1−6 These statements triggered interest in the incorporation of PUFA into processed food products. However, challenges in supplementing processed foods with omega-3 fatty acids include both their susceptibility to oxidation given the high concentration of unsaturation, promoting free radical initiation, and their physical incompatibility with dry or aqueous ingredients.7 These technical issues can be addressed through emulsification and encapsulation with appropriate dry carriers, thus increasing the shelf life of food matrices enriched with n-3 polyunsaturated fatty acids. Emulsification can be achieved via the preparation of oil-inwater emulsions containing small spherical oil droplets that are stabilized in the aqueous phase by proteins and surface active polysaccharides including arabic gum and modified starch.8,9 These surface-active hydrocolloids are adsorbed at the interface between the oil and aqueous phase to prevent oil droplets from aggregation by lowering surface tension and increasing force of repulsion and thus minimize oil oxidation. The use and the selection of biopolymers blends as emulsifier may allow increasing the emulsification efficiency and its stability.10−12 Proteins such as whey protein and gelatin have been used to form oil-in-water (o/w) emulsions and improve the physicochemical stability of PUFA.8,9,13−15 Other authors have stated that whey protein isolate could adsorb rapidly at the Published XXXX by the American Chemical Society

oil−water interface and provide structural support for oil droplets through a combination of electrostatic and steric interactions.16,17 Fish gelatin has been also widely used in food and pharmaceutical industries due to its unique functional and technological properties.18,19 This protein is a relatively high molecular weight protein, which allows the creation of o/w emulsions with positively charged droplets.13,20 Gelatin is produced in two types depending upon the use of acid (type A, pI ∼7−9) or alkaline (type B, pI ∼5) for extraction.21 The combination of both proteins could be promising for the prevention of oxidative deterioration in PUFA.9 The layer-by-layer (LBL) polyelectrolyte deposition technique has gained a great deal of popularity for preparing polyelectrolyte microcapsules because of its ability to create highly tailored microcapsule shells of various sizes.22 To prepare these microcapsules of different size, the assembly process is driven by the electrostatic attraction of oppositely charged materials (polyanions and polycations) to form the microcapsule shells.23,24 These microcapsules are obtained via stepwise addition of positively and negatively charged polyelectrolytes such as hydrocolloids and proteins onto colloidal particles.25 The properties and functionalities of these microcapsules such as the encapsulation efficiency and release characteristics as well as the protection against oil oxidation can be modulated by varying the microcapsule Received: July 7, 2015 Revised: October 7, 2015 Accepted: October 12, 2015

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DOI: 10.1021/acs.jafc.5b00858 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Journal of Agricultural and Food Chemistry

Article

Table 1. Combination Used for the Preparation of Secondary Multilayer Emulsions emulsion primary emulsion secondary emulsions primary emulsion secondary emulsions

1

2

3

0.5% WPI + 0.75% FG

0.5% FG + 0.75% WPI

After homogenization, these individual primary emulsions were used to prepare secondary multilayer emulsions to form (a) anionic− cationic emulsion by mixing WPI primary-based emulsion with the aqueous gelatin protein solution in a ratio of 1:1 and (b) cationic− anionic emulsion by mixing FG primary-based emulsion with the aqueous whey protein solution (ratio 1:1 at pH 6.5) previously prepared at various concentrations of WPI or FG. The final ratios of the secondary emulsions are described in Table 1. These secondary emulsions containing a final oil concentration of 5% were then passed through a high-pressure homogenizer for two passes at 500 psi to avoid bridging flocculation. Once completed, these emulsions were stored at 21 °C for the daily analysis. Droplet Size Measurements. Droplet size distributions were determined after preparation of the primary and secondary emulsions using the granulometer Mastersizer 2000 laser light scattering instrument (Malvern Instruments Ltd., Worcestershire, UK) equipped with a Hydro 2000 mV handling unit (containing the buffer solution). The samples were stirred continuously within the sample cell to ensure homogeneity at ambient temperature. Droplet size distributions were calculated by the instrument according to the Mie theory, which uses the refractive index difference between the droplets and the dispersing medium to predict the intensity of the scattered light. The ratio of the refractive index of flaxseed oil (1.48) to that of dispersion medium was 1.11. Droplet size measurements were reported as both

thickness via wall thickness and composition. The average electric charge of the microcapsules can be controlled via the acidity adjustment of the continuous phase, the variation of the number of layers, or changing the characteristics of the polymers,24 thus offering the possibility to control the interactions between the charged polymers. To the best of our knowledge, the microencapsulation of flaxseed oil using the LBL technique with different types of wall materials and the study of its impact on the oxidative stability of flaxseed oil have not been reported in the literature. The LBL assembly technique to produce microcapsules using flaxseed o/ w emulsion droplets and food-grade protein as wall materials, at different concentrations to modulate the size distribution of the emulsion droplets, could be an interesting approach to enhance the physicochemical stability of the flaxseed oil. The objective of this study was to investigate the effects of concentration of whey protein isolate (polyanion) or fish gelatin (polycation) materials as alternate first layer to produce microcapsules at pH 6.5 via the LBL self-assembly process and to study the physicochemical properties of the microcapsules and their ability to protect the flaxseed oil against oxidation over storage time.

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Batch 1: Anionic−Cationic Multilayer Emulsion 1% WPI + 10% flaxseed oil 0.5% WPI + 0% FG 0.5% WPI + 0.1% FG 0.5% WPI + 0.5% FG Batch 2: Cationic−Anionic Multilayer Emulsion 1% FG + 10% flaxseed oil 0.5% FG + 0% WPI 0.5% FG + 0.1% WPI 0.5% FG + 0.5% WPI

(a) volume-length mean diameter (d43)

MATERIALS AND METHODS

d43 =

Materials. Fish gelatin (275FG 30) type A, 270 gel strength, 88% protein, 20 mesh, was provided by Rousselot Inc. (Mukwonago, WI, USA). Powdered whey protein isolate (Hilmar 9400) was obtained from Hilmar Ingredients (Hilmar, CA, USA). As stated by the manufacturer, the protein content was 94% (dry basis), and lactose and ash contents were 0.2 and 2.0%, respectively. Analytical grade sodium phosphate monobasic monohydrate (NaH2PO4·H2O), sodium phosphate dibasic heptahydrate (Na2HPO4·7H2O), and hexanal (volatile) were purchased from Sigma Chemical Co. (St. Louis, MO, USA). Deionized water was used for the preparation of all emulsions. Virgin cold-pressed organic flax oil was acquired from Maison Orphée (Quebec, PQ, Canada). The claim by Maison Orphée was that the flax oil contained 53.0% omega-3 fatty acids. Preparation of Emulsions. A pH 6.5, 50 mM stock buffer solution was prepared using 0.5 M NaH2PO4 and 0.5 M Na2HPO4 in deionized water. Sodium azide (0.05%) was added to the buffer solution to ensure microbial stability of the emulsions. The stock buffer solution was used to prepare the emulsifier solutions containing 1% whey protein isolate (WPI) and 1% fish gelatin (FG). To ensure proper dispersion, both emulsifier solutions were mixed separately for 2 h at 37 °C and then left overnight at 37 °C for complete hydration. Emulsifier solutions and the oil phase were tempered at room temperature prior to the preparation of the emulsions. Preparation of Anionic−Cationic and Cationic−Anionic Multilayer Emulsions. The primary o/w emulsions were prepared by emulsifying 10% flaxseed oil with protein solutions containing 1% concentration of WPI or FG hydrated in the buffer solutions at room temperature, using a Polytron PT-35 at 3500 rpm for 3 min followed by three passes at 5000 psi and one pass at 500 psi in a high-pressure valve homogenizer (Emulsiflex C-5, Avestin, ON, Canada). The emulsions were collected in a beaker and submerged with an ice/water mixture to maintain the ambient temperature as constant as possible.

∑ ni × di 4/∑ ni × di 3

(b) surface-weighted mean diameter

d32 =

∑ ni × di 3/∑ ni × di 2

where ni is the number of droplets of diameter di. The particle polydispersity could be given by two factors/values, namely, uniformity (how symmetrical the distribution is around the median point)

uniformity =

∑

Xi[d(v , 0.5) − di] D(v , 0.5)ΣXi

where Xi is the total number of particles and the span (the width of the distribution) is defined by the expression

span = (D90 − D10)/D50 D90 is the drop diameter below which 90% of the distribution exists, D50 is the median diameter, and D10 is the drop diameter below which 10% of the distribution exists. The smaller the span value the narrower the particle size distribution. Surface Net Charge Measurements. Measurements of the electrical charge (zeta-potential) of the lipid droplets were evaluated by electrophoresis and dynamic light scattering measurements (Zetasizer Nano-ZS, Malvern Instruments) immediately after emulsification. The emulsions were diluted 1:300 in the same phosphate buffer to avoid multiple scattering effects. The viscosity of this diluted emulsion was measured at a shear rate of 0.1/s (viscosity at rest), and the value was entered into the Zetasiser software. Each individual ζ-potential data point was calculated from the average of at least six readings made on the duplicate samples. B



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Figure 1. Illustrative Turbiscan backscattering profile of a typical instability phenomenon encountered in emulsions over time. Each curve corresponds to a single time measurement. Reprinted from ref 29. Protein Load at the Oil−Water Interface. From each freshly made emulsion, a weighted quantity was centrifuged at 25200g for 1 h at 4 °C using the Beckman centrifuge model J2.21 and rotor JA-20.1 (Beckman Instruments, Fullerton, CA, USA) to accelerate the migration of the dispersed particles. Protein assays were done with the serum according to the Kjeldahl method (N × 6.38) (Kjeltec Auto 1030 Analyzer, Foss Analytical, Sweden). By subtracting the amount of protein in serum from the total amount in the emulsion formulation, respective concentrations in the creamed layer were determined as weight percent units. Those values were then divided by the respective interfacial area estimated from initial droplet diameter (d32), allowing evaluation of interfacial protein load (mg/m2).26 All measurements were done in duplicate for each emulsion. Gel Electrophoresis. Sodium dodecyl sulfate−polyacrylamide gel electrophoresis (SDS-PAGE) was carried out according to the method of Laemmli27 on Bio-Rad (St. Louis, MO, USA) precast 10−20% TrisHCl gel, using the Bio-Rad Criterion cell. SDS-PAGE was used to identify the protein subunits present in dried cream. The cream layer of each emulsion was separated by centrifugation at 100000g for 40 min at 4 °C using a Beckman centrifuge (Beckman model J2.21) and a rotor model JA-20.1 (Beckman Instruments). The separated cream was transferred into another tube, dispersed well in demineralized water, and centrifuged at 87000g for 30 min at 4 °C. The washing procedure was repeated twice, and the resulting cream layer was freeze-dried prior to analysis. Samples (2.5 mg/mL) were dispersed in Laemmli sample buffer (Bio-Rad) plus 5% β-mercaptoethanol and vortexed for 15 min. The dispersions were heated at 100 °C for 5 min and centrifuged before loading. Twenty microliters of sample solution was loaded in each well and the gel run at a constant voltage of 150 V. The gel was stained with Coomassie brilliant blue R-250. Bio-Rad Precision Plus Protein Standards (10−250 kDa) were used as molecular markers. Rheological Properties. Measurement of rheological parameters was based on the methods by Taherian et al.28 using an AR1000 rheometer (TA Instrument, New Castle, DE, USA) equipped with a 2° cone of 60 mm diameter. Emulsions were subjected to flow test, measuring shear viscosity (ηγ) as a function of shear rate ranging from 0.1/s to 100/s at 22 °C. Flow behavior index (n) and consistency coefficient (m) were calculated using the power law model. Using flow behavior index (n) and consistency coefficient (m), the apparent viscosity at 0.1/s (viscosity at rest) for each emulsion was calculated and the outcomes were compared. The duplicate samples along with three measurements were considered. Emulsion Stability. The destabilization of the emulsions was evaluated using a Turbiscan (Quick scan, Coulter Corp., Miami, FL, USA). The sample (6 mL) was placed in a cylindrical glass cell (100 mm height, 16 mm internal diameter) and scanned from the bottom to the top with a light source (λ = 850 nm). Simultaneously, two

synchronized optical sensors respectively recorded light transmitted through the sample (180° from the incident light) and light backscattered by the sample (45° from the incident radiation). These data are represented in curves of transmittance (%T) and/or backscattering (%BS) as a function of the sample height (mm) (Figure 1).29 The scans are repeated over time, each time providing a single curve, and at the end of the experiment, all curves are superimposed on the same graph to show the overall destabilization of the system. This complete analysis mode enables the detection of both the migration phenomena and the variation of average particle size. To better visualize the changes undergone in the systems, it is convenient to work in the reference mode (Δ%BS or Δ%T), which subtracts the first curve (t = 0) from the subsequent ones to see the variations of profiles related to the initial state. The profiles recorded for each sample were then analyzed and expressed either (i) as the average Δ% BS variation measured at the early zone of the profiles (5−10 mm) corresponding to the clarification zone; (ii) the middle zone of the profiles (30−40 mm) as a function of time, which is related to the variation of droplet size; and (iii) the end of the profiles, which corresponds to the creaming zone. Our measurements were performed immediately after preparation of emulsions and at different times. The curves obtained by subtracting the BS profile at t (ΔBS = BSt − BS0) display a typical shape that allows a better visualization of the destabilization processes. In addition, 100 mL of each emulsion sample was placed in a Wainthropp tube to observe the visual separation. Assessment of Oxidative Stability. Measurement of Hexanal Concentration. A volatile secondary oxidation compounds (hexanal) was selected as an indicator for flaxseed oil oxidation and was extracted from emulsions by headspace solid-phase microextraction (HSSPME). Emulsions were stored in 10 mL vials, sealed with a magnetic screw cap containing a polytetrafluoroethylene (PTFE)/silicone septum (Varian, Mississauga, ON, Canada) at 21 °C and exposed to two fluorescent lights located 30 cm above the emulsion. At 1, 3, 7, 10, and 21 day intervals, each vial containing the emulsion sample was transferred for volatile analysis. The HS-SPME fiber (85 mm Carboxen/PDMS, Supelco, Oakville, ON, Canada) was inserted into the headspace of the vial for 44 min at 40 °C. The SPME operations were automated using an MPS2 multipurpose sampler (Gerstel Inc., Baltimore, MD, USA). Volatile compounds were desorbed by inserting the fiber into the injection port 1078/1079 of a Varian CP-3800 gas chromatograph (Palo Alto, CA, USA) in splitless mode (glass insert SPME, 0.8 ID; Varian, Mississauga, ON, Canada) for 3 min at 300 °C. The GC-MS system used in this study consisted of an ion trap mass spectrometer equipped with an electronic impact ionization source controlled with Saturn 2000 mass spectrometry detector (Varian Inc., Palo Alto, CA, USA). A Varian CP-Sil 8CB-MS capillary column (5% phenyl/95% dimethylpolysiloxane; 30 × 0.25 mm; 25 μm film C



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Table 2. Average Particle Sizes (d3,2) and (d4,3) for Primary and Secondary Emulsionsa emulsion primary emulsion 1% WPI secondary emulsions 0.5% WPI + 0% FG 0.5% WPI + 0.1% FG 0.5% WPI + 0.5% FG 0.5% WPI + 0.75% FG primary emulsion 1% FG secondary emulsions 0.5% FG + 0% WPI 0.5% FG + 0.1% WPI 0.5% FG + 0.5% WPI 0.5% FG + 0.75% WPI

d43 (μm)

d32 (μm)

spanb

uniformity

0.62 ± 0.015

0.45 ± 0.124

1.59

0.467

± ± ± ±

1.61 1.66 1.60 1.65

0.471 0.491 0.481 0.491

1.05 ± 0.01

11.91

3.02

± ± ± ±

1.92 1.91 1.81 1.87

0.682 0.672 0.683 0.672

0.53 1.08 1.26 1.58

± ± ± ±

0.31c 0.01b 0.06ab 0.10a

0.59 0.66 0.80 0.97

4.31 ± 0.11 13.91 3.69 2.99 2.50

± ± ± ±

1.17a 0.00b 0.06b 0.22b

2.06 2.21 1.61 1.50

0.18b 0.00b 0.01ab 0.02a

0.31a 0.01a 0.01b 0.01bc

Values are means ± SD for n = 3. Means within the same column followed by different letters are significantly different (P < 0.05). bRelative span factor (RSF): a dimensionless parameter indicative of the uniformity of the drop size distribution.

a

thickness) was used with helium as carrier gas at a flow rate of 1.0 mL/ min. The column oven was set initially at 35 °C for 3 min, heated to 80 °C at a rate of 6 °C/min, increased to 280 °C at a rate of 20 °C/ min, and then held at 280 °C for 2 min. The total time of analysis was 22.5 min. The mass spectrometer was operated in the mass range from 30 to 200 at a scan rate of 1.00 s/scan. Calibration curves were done using standards of hexanal in media containing either 0.5% WPI or 0.5% FG at pH 6.5. Hexanal retention time was around 6.4 min. The quantification was realized by total ion current mode. Measurement of Conjugated Dienes Hydroperoxide (CD). A modification of the method described by IUPAC 2.50530 has been used for the quantification of the conjugated dienes hydroperoxides over storage period. The emulsion sample (200 μL), which was stored under the same condition for hexanal analysis, was added to a mixture of 10 mL of isooctane/2 propanol (2:1 v/v) to obtain an optical density value (OD) between 0.2 and 0.8 and vortexed 30 s. The absorbance was measured at 232 nm using an UV−vis scanning spectrophotometer (Cary 300 BIO UV/visible). For the protein-based emulsions, a filtration through a 0.45 μm PUDF Nagel filters (Canadian Life Science, Peterborough, ON, Canada) was applied prior to the measurement to remove protein from the sample and thus reduce its spectrum interference in this region. Measurements of the CD were performed in triplicate on each emulsion, and averaged data were expressed in millimolar CD per kilogram of oil using 27000 cm−1 as the molar extinction coefficient. Statistical Analysis. Experimental analyses were repeated twice, and each measurement was made at least in triplicate. The results were averaged (n = 6) and statistically analyzed by one-way analysis of variance (ANOVA) using PRISM software, version 3.02 (Graph Pad Software, Inc. San Diego, CA, USA). Significant differences between means were determined by Tukey’s multiple-comparison test procedure at the 5% significance level.

often produces relatively large droplet sizes during homogenization,32 with larger span and polydispersibility values compared to WPI.33,34 Dilution of primary emulsions from 1% to form 0.5% with no addition of complementary protein (control secondary emulsions) followed by gentle homogenization pressure at 500 psi did not affect significantly the particle size in the case of WPI (0.5% WPI + 0% FG), whereas a significant increase in the droplet particle size (13.91 μm) was observed in the case of FG (0.5% FG + 0% WPI). These large particles may result from droplet aggregation and/or flocculation. Indeed, the span values as shown in Table 2 representing the particle polydispersibility confirmed the high span value for FG and therefore the large particle size distribution of FG primary emulsion. Another possible explanation could be the fact that the low emulsifying capacity of FG seems to be explained by its reduced mobility, disordered structure, and possibly a homogeneous distribution of hydrophobic residues (low amphiphilicity). The viscosity of hydrated FG, at the identical concentration, was significantly (P < 0.05) higher than that of hydrated WPI (as shown further in Figure 4). For the anionic−cationic emulsions, the deposition of FG over the WPI as anionic polymer at concentrations from 0.1 to 0.75% increased significantly (P < 0.05) the droplet size of the secondary emulsion from 1.08 to 1.58 μm (d43) and from 0.66 to 0.97 μm (d32). On the other hand, for the cationic−anionic emulsion in which the gelatin is used as the primary emulsion forming the first absorbing layer, the addition of WPI varying from 0.1 to 0.75% decreased significantly (P < 0.05) the droplet size from 3.69 to 2.50 μm (d43) and from 2.21 to 1.50 (d32). The other interesting observation is that the mean oil droplet size for FG primary emulsion is reduced after deposing WPI (secondary emulsions) with increasing concentration of WPI. This effect is opposite for those made with WPI as primary emulsions because raising the concentration of FG had for effect an increase in the volume and mean surface diameter. There are many possible reasons to account for the observed decrease in mean droplet size after deposing WPI over FG added primary emulsion such as (a) the interface creation stabilized by the addition WPI resulted in the breakage of aggregates, yielding higher tensioactive properties; (b) the second low-pressure homogenization treatment resulted in an increase in total droplet surface area that could be stabilized by

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RESULTS AND DISCUSSION Particle Size Measurements. The mean average droplet size for primary and secondary emulsions for both the anionic− cationic and cationic−anionic multilayer emulsions is provided in Table 2. The data revealed that the primary emulsion prepared with 1% WPI had lower droplet size than emulsion prepared with 1% FG, suggesting that WPI, having high tensioactive properties, reached easily the interface of oil droplets, causing further breakdown of the droplets during prehomogenization and homogenization processes. Some previous studies have shown that gelatin is less surface-active, but is still capable of acting efficiently as an emulsifier in o/w emulsions.20,31 Nevertheless, when used on its own, gelatin D



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the protein and a dislocation of aggregates; and (c) the frequency of droplet collisions decreased due to the increase in aqueous phase viscosity. All of the above factors could facilitate droplet disruption and prevent droplet coalescence within the homogenizer, thus forming smaller droplet sizes.14,23 Particle Charge. Droplet charge was quantified at pH 6.5, which is above the isoelectric point of WPI (pI ≈4.6−5.6) and slightly lower than that of FG (pI ≈7−9), respectively. The net negative charge intensity for WPI primary emulsion was −24.6 mV, whereas FG primary emulsion droplets showed a positive net charge, but close to 0 (Table 3). The electric charges of Table 3. Comparison of ζ-Potential for Primary and Secondary Emulsions

Figure 2. SDS-PAGE patterns of whey protein isolate (WPI), fish gelatin (FG), and WPI+FG mixture as control protein solutions and of the creams of primary emulsions (P1−P3) and secondary emulsions (S1−S8) after separation and centrifugation. Samples were run on 10− 20% Tris-HCl gel. Lanes: Std, molecular weight markers; P1, 1% WPI; P2, 0.5% WPI; P3, 0.5% FG; S1, 0.5% WPI; S2, 0.5% WPI + 0.1% FG; S3, 0.5% WPI + 0.5% FG; S4, 0.5% WPI + 0.75% FG; S5, 0.5% FG; S6, 0.5% FG + 0.1% WPI; S7, 0.5% FG + 0.5% WPI; S8, 0.5% FG + 0.75% WPI. All emulsions were prepared at 5% oil content.

ζ-potential (mV) emulsion

0.5% 0.5% 0.5% 0.5%

WPI WPI WPI WPI

+ + + +

FG FG FG FG

secondary

−23.91 −21.75 −14.30 −11.15

0% FG 0.1% FG 0.5% FG 0.75% FG

± ± ± ±

0.91a 1.08b 0.17c 0.12d

presence of low molecular weight bands of β-1actoglobulin ∼20 kDa and α-lactalbumin ∼15 kDa, appearing at the bottom of the gel, as well as the presence of high molecular weight proteins, which occupy the large fractions of fish gelatin, are observed on the top of the gels. Protein solution prepared with WPI−FG mixture showed a predominance of WPI polypeptide subunits, whereas the high molecular weight proteins related to FG and its dimer look very faint. Similar gel quality was observed by Taylor et al.37 after the addition of whey protein isolate into the gelatin sample. For WPI, FG, and WPI + FG used as primary and/or secondary emulsions, the pattern of adsorbed proteins indicated that surface proteins are mostly related to WPI and proteins related to FG are either absent or present in only small amounts. This may indicate that addition of WPI to FG contributed to the formation of a new WPI−FG protein complex that is incorporated into the interface of the oil droplets, contrarily to the case in which FG is incorporated to WPI. Similar patterns were observed by Taherian et al.9 using a combination of WPI and FG for emulsion preparation. These authors suggested that pH and its intimacy to the isoelectric point of protein play a major role in the amount of protein adsorbed to the surface of oil droplets. Moreover, amphiphilic WPI structures are more flexible compared to those of fish gelatin as stated by Jiang et al.38 This could cause faster adsorption and prevalence of WPI proteins at the surfaces of oil droplets. The above results were confirmed by assessing the protein load as presented in Figure 3. As illustrated, the amount of protein adsorbed onto the fat surface (protein load) for emulsions prepared by FG are significantly lower compared to WPI-based emulsions, demonstrating the high surface active properties of WPI proteins over FG. In addition, increasing the concentration of FG protein used to yield secondary emulsions did not show a significant difference in the amount of adsorbed protein. Interestingly, the results of zeta potential suggested the evidence of FG deposing over WPI. However, the adsorption strength was weaker, likely due to the harsh centrifugation conditions and the washing steps to recover the cream layer. These conditions seem to release the FG layer. Unlike the WPI

+1.09 ± 0.04

1% FG 0.5% 0.5% 0.5% 0.5%

primary −24.60 ± 1.06

1% WPI

+ + + +

0% WPI 0.1% WPI 0.5% WPI 0.75% WPI

+0.71 −2.48 −3.28 −3.66

± ± ± ±

0.05a 0.18b 0.21c 0.07d

WPI- and FG-coated droplets are controlled by the ionization degree of amino groups (−NH2) and carboxyl groups (−COOH) of protein molecules, which are dependent on the pH of the surrounding aqueous phase. Because the emulsion pH is close to the isoelectric point of FG, the droplet’s charge is close to zero. In other words, in FG-coated emulsions, the number of positively charged groups is close to the number of negatively charged groups and the net surface charge of the droplets becomes almost neutral.35 Deposition of FG over WPI as primary emulsion at concentration of 0.1− 0.75% decreased significantly (P < 0.05) droplets’ charge in the secondary emulsions (from −21.75 to −11.15 mV) at pH 6.5. On the other hand, the secondary emulsions made by deposing WPI over FG primary emulsion at the same concentration raised slightly the net electronegativity of the multilayered emulsion (from −2.48 to −3.66 mV). The net charge and low negative charge intensities of primary or secondary emulsions coated with FG are due to the paired positive and negative charges and proximity of FG’s isoelectric point to the pH of the system.9,36 It is important to note that by looking closely at the secondary emulsions prepared at the same concentration (0.5% WPI and 0.5% FG), the order of deposition affects differently the net surface charge of the oil droplets. The effect of protein deposition on particle charge was revealed by the change in zeta potential following protein deposition. For fish gelatin, the zeta potential change was 10.3 mV (|−24.6 to −14.3|), whereas for WPI it was 4.4 mV (|−3.28 to 1.09|). SDS-PAGE and Protein Load. The SDS-PAGE patterns of individual, mixed protein solutions as well as adsorbed proteins in emulsions prepared with WPI, FG, and as mixture (multilayer emulsions) in buffer at pH 6.5 are illustrated in Figure 2. As a control protein solution, WPI revealed the E



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Figure 3. Protein load of primary and secondary emulsions containing various amounts of WPI and/or FG prepared with 5% oil.

Figure 4. Apparent viscosities (γ = 0.1/s) of primary and secondary emulsions containing various amounts of WPI and/or FG prepared with 5% oil.

used to form the primary emulsion, adsorption of the secondary layer FG seems to involve weak interaction links. On the other hand, addition of WPI to primary FG-based emulsion at various concentrations from 0.1 to 0.75% was accompanied by significant (P < 0.05) adsorption of this protein at the surface of the gelatin primary emulsions, increasing therefore the protein load by 2−3-fold, respectively (Figure 3). It is important to note that in the case of the FG−WPI system, whether the observed increase in protein load values could be related to a real layer-by-layer deposition of whey

protein above FG or rearrangement of WPI−FG protein complex at the surface of oil droplets remains unknown and requires further investigations. Apparent Viscosity of Emulsions. The apparent viscosity of prepared emulsions is presented in Figure 4. For primary emulsions prepared at 1% WPI or 1% FG with 10% oil content, FG emulsion showed higher viscosity than WPI emulsion at the same concentration. Damodaran et al.39 reported that the denatured dispersions of randomly coiled molecules (e.g., fish gelatin) show greater viscosity than solutions of compact folded F



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Figure 5. Variation in backscattering (Δ%BS) in the 10 mm zone monitored over 21 days for (A) whey protein primary coated and (B) fish gelatin primary coated emulsions at different concentrations.

droplets of WPI−FG-stabilized emulsion decreased markedly (became less negative), along with a significant increase in apparent viscosity. In other words, deposition of a lower amount gelatin on whey protein coated droplets decreased the zeta potential and increased the apparent viscosity significantly (P < 0.05). Similarly, higher viscosity was also associated with deposition of WPI at different concentrations over FG as primary emulsion, which also correlated positively with the significant increase in protein load. Interestingly, independent of the deposition order, secondary emulsions prepared at the same concentrations (0.5% WPI and 0.5% FG) showed similar apparent viscosities (Figure 4), whereas for the other physicochemical parameters, WPI-based emulsions showed lower particle size, higher electronegativity, and greater protein load. Physicochemical Stabilities of Emulsions. The degrees of emulsion stability for WPI and FG alone and in

globular molecules (e.g., WPI). With WPI primary emulsion, the increase in FG concentration from 0.1 to 0.75% was associated with a significant (P < 0.05) increase in the apparent viscosity (0.1/s) of secondary emulsions (from 1.92 to 3.96 mPa·s). This increase in the apparent viscosity could be attributed to electrostatic association between the two polymers as well as the interaction of water−gelatin present in the aqueous phase. Several authors reported that associative interactions involve electrostatic attraction between polyanion and polycation protein polymers, resulting in an increase in apparent viscosity.40 Similarly, addition of WPI at different concentrations to FG primary emulsion resulted in a gradual increase in apparent viscosity, but to a lower extent. Considering the protein load (Figure 3), the viscosity (Figure 4), and the zeta potential (Table 3), the results showed that in the case of WPI, addition of FG from 0 to 0.75% did not raise the protein load significantly, whereas the zeta potential of the G



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Figure 6. Evolution of conjugated dienes for (A) whey protein primary coated and (B) fish gelatin primary coated emulsions.

combinations at pH 6.5 are shown in Figure 5. The figure shows the decrease in backscattering at a fixed height (10 mm), during the studied time, which can be related to the kinetics of migration of small particles as a result of emulsion destabilization. As was theoretically demonstrated,41 the intensity of backscattered light is related to the number of droplets located at a specific height of the emulsion. Therefore, the higher the decrease in BS intensity, the higher the destabilization of the emulsion related to the migration of particles. In our case, the stability of emulsions by analyzing the BS profiles at different storage times revealed that compared to FG, primary emulsion prepared with WPI (0.5 or 1% concentration) had high stability, due to both lower particle size and its amphiphilic character, which provided good protection to the oil droplets through a combination of electrostatic and steric interactions, stabilizing the emulsions against emulsion destabilization.8 Addition of FG from 0.1 to 0.75% resulted in a significant reduction of Δ%BS, indicating greater destabilization of the emulsions. IAn evident decrease in BS at the bottom of the tube and a concomitant increase of BS in the upper zone (data not shown) attributed to the formation of a cream layer were noted from the quickscan profiles. The top of the profile had a shape indicative of the migration of individual particles.

Furthermore, at higher concentration of FG (0.5 and 0.75%), WPI−FG secondary emulsions resulted in the appearance of a slight oil ring at the top of the Wainthropp tube after 1 month of storage. The phase lifting observed in the conical tube may result from droplet coalescence. When the concentration of FG increases, the complexes between FG and WPI are formed in the aqueous phase and gradually lose the ability to “bridge” the oil droplets. Higher concentrations of FG may have destabilized the WPI primary emulsion, which favored more the formation of FG−WPI protein aggregates rather than creating a second layer around the WPI primary emulsion droplets for higher stability. As stated earlier, for primary emulsion prepared with 1% FG, both creaming and flocculation mechanisms were involved. The quickscan profile showed that the creaming process occurred during the first 24 h. The profile had a bell-shaped curve at the bottom of the tube, typical of migration of particles. Then, there was a great increase in BS at the top of the tube, which suggested migration of aggregates. Addition of a low concentration of WPI (0.1%) to FG primary emulsion exhibited a decrease of Δ%BS. However, addition of 0.5 and 0.75% WPI resulted in a re-increase in Δ%BS, suggesting the formation of lower droplet size, which greatly promoted emulsions’ physical stability compared to FG alone. This H



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Figure 7. Influence of lipid oxidation on hexanal formation by HS-SPME and GC-MS using CAR/PDMS fiber during 21 days of storage under UV light.

complex phenomenon, which includes the adsorption, desorption, unfolding, and aggregation, occurring simultaneously at the interface.42 Pearce and Kinsella43 stated that the ability of the protein to be adsorbed at the surface of oil droplets depends upon its capacity to unfold and spread over the interface to stabilize the new area created. This adsorption process can be easily affected by various factors such as the structure of the protein, the protein hydrophobicity, and the chemical environment of the solution such as the pH and ionic strength. The adsorption process can also be affected by the physical state of the protein such as concentration and temperature, which can cause conformational change or partial denaturation of the adsorbed protein layer at the oil/water interface.44 The structure−function properties of adsorbed protein at the interface affect the stability of a prepared emulsion.45,46 Formation of CD Hydroperoxides and Hexanal. The present work attempts to relate the impact of the anionic− cationic versus cationic−anionic LBL deposition technique on

improvement in stability could be explained by the fact that at 0.1% WPI, FG is displaced by WPI, forming a fine emulsion. The second homogenization treatment may have possibly affected the rearrangement of protein at the surface of oil droplets, enhancing faster spreading of whey proteins at emulsion droplets’ surface. In addition, we must also consider the increase in viscosity, which can slow the phase separation mechanism. Thus, higher destabilization was observed specifically when FG was added with increasing concentration as a multilayer polymer on WPI primary emulsions. In parallel, the increase in d2,3 values (Table 2) reflected the observed destabilization process. Interestingly, as shown in Figure 5, destabilization occurred to lesser extent as WPI concentration added to FG primary emulsions increased. Indeed, the decrease in droplet size with the addition of WPI to FG primary emulsions resulted from the high tension-active property of WPI; hence, a greater interfacial protein concentration will promote a higher physical stability of these emulsions. The stability of an emulsion is a I



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0.5% WPI + 0.5% or 0.75% FG, the presence of WPI at the interface and FG in the bulk may prevent the binding of some pro-oxidant impurities (such as transition metals) and protect flaxseed oil droplet against oil oxidation. In summary, the stability of multilayer emulsions prepared using two protein polymers (WPI and FG) was dependent on both the order used to assemble the biopolymer layers and the concentration of biopolymers. Primary emulsions prepared with 0.5% WPI had lower particle size droplets than emulsions prepared with 0.5% FG. The deposition of FG at different concentrations over the WPI primary emulsion increased the droplet particle size significantly, whereas WPI deposited over FG decreased it. Addition of FG reduced the electronegativity and WPI increased it. Similarly, coating gelatin over WPI caused a synergistic increase in the apparent viscosity, but to a lower extent when WPI was used as secondary emulsion layer. Electrophoresis showed that for WPI, FG, and WPI + FG used as primary and/or secondary emulsions, the pattern of adsorbed proteins indicated that surface proteins are mostly related to WPI, and proteins related to FG are either absent or in very low amount. Thus, WPI was presented in higher concentration at the interface compare to FG as confirmed by protein load results. From these results, it is also clear that the stability of WPI emulsions was strongly affected by the addition and concentration of fish gelatin protein rather than addition of WPI over FG. The design of multilayer emulsions with WPI first having good interfacial properties coated with a second oppositely charged biopolymer (protein or polysaccharide) could have important implications for the formulation of many types of commercially important emulsion-based products.

the emulsion sensitivity to oxidation. Chemical stability is described by the degree of lipid oxidation, which is a complex sequence of chemical changes that result from the interaction of lipids with oxygen-active species. The most likely mechanism for the acceleration of lipid oxidation or chemical instability in emulsions is the decomposition of lipid hydroperoxides (ROOH) into highly reactive peroxyl (ROO) and alkoxyl (RO) radicals by transition metals or other pro-oxidants.47 The oxidative status of each emulsion stabilized with proteins has been expressed as an oxidative factor based on the change of CD at 232 nm as primary oxidation byproducts and hexanal as secondary byproduct over storage under accelerated conditions. Obviously, higher CD and hexanal values reflect the progress of oxidative deterioration in these emulsions. As shown in Figure 6, a rapid production of CD was observed within the first 20 h with all of the emulsions, and then the values tend to plateau due to the conversion of dienes to other byproducts. In terms of CD production, the results indicated that WPI and FG primary emulsions showed similar oxidation deteriorations, which generated almost an equal amount of CD of 24 mmol/kg oil after 20 h to reach later a final concentration of 34 mmol/kg oil after 60 h. Addition of FG at 0.1, 0.5, and 0.75% concentrations to WPI primary emulsions markedly decreased the CD production by 11.5, 27.4, and 29.5%, respectively. Whether the reported increase in the emulsion droplet size (as reported in Table 2) may explain this oxidative resistance is not certain. The literature evidence concerning the influence of oil droplet size and interfacial area on lipid oxidation of emulsions is variable and generally contradictory. In recent works, Kiokias et al.48 and Dimakou et al.49 reported no clear effect of droplet size on the oxidative destabilization of whey protein emulsions, in terms of both CD and thiobarbituric acid-reactive substance (TBARs) oxidative indicators. Contrarily, addition of WPI at 0.1 and 0.5% to FG primary emulsions increased the CD production by 9.34 and 7.14%, respectively, whereas with 0.75% WPI the CD decreased by 6.54% above 20 h of storage. Due to its amphiphilic character, WPI may have offered a stronger barrier to prooxidant agents present in the aqueous phase, preventing them from readily approaching the lipid core of the emulsion droplets. Such an effect may account for the increased oxidative stability of emulsions prepared under the present experimental conditions. Changes in hexanal concentration were monitored during emulsion storage (Figure 7). The results confirmed the tendency observed for the CD generation; thus, an increase of FG concentration deposited over the WPI was associated with a decrease of oxidative sensitivity, and the lowest extent of hexanal was noted with 0.5% WPI + 0.75% FG (0.32 ppm) versus 0.50 ppm of hexanal with the counterpart emulsion prepared with 0.5% FG + 0.75% WPI. Interestingly, emulsions prepared at the same concentrations of WPI and FG exhibited different oxidative protection. Deposition of fish gelatin on whey protein coated droplets (0.5% WPI + 0.5% FG) showed a lower hexanal value than the hexanal content observed for WPI added to fish gelatin coated droplets. Thus, whey protein showed higher ability than fish gelatin to reduce lipid oxidation. Literature reports that the extent of lipid oxidation is related to the thickness of the protein layer at the interfacial region of emulsion droplet, affecting interactions between lipids and aqueous phase prooxidant and chemical components of proteins that enable scavenging of free radicals or chelating of pro-oxidant metals.50 Therefore, in emulsions stabilized by

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AUTHOR INFORMATION

Corresponding Author

*(P.F.) Phone: (450) 768-3256. Fax: (450) 773-8461. E-mail: [email protected]. Funding

This project was funded by Agriculture & Agri-Food Canada under the growing forward health claims, novel foods and ingredients initiative. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We thank Marc André Nadeau (Sherbrooke University) for his assistance with the preparation of the emulsions and analyses conducted during this study.

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REFERENCES

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