pectin-stabilized multilayer emulsions as astaxanthin delivery systems

International Journal of Biological Macromolecules 140 (2019) 985–997

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International Journal of Biological Macromolecules journal homepage: http://www.elsevier.com/locate/ijbiomac

Formation, characterization, and application of chitosan/ pectin-stabilized multilayer emulsions as astaxanthin delivery systems Chengzhen Liu a, Yunbing Tan b, Ying Xu a, David Julian McCleiments b,⁎, Dongfeng Wang a,⁎⁎ a b

College of Food Science and Engineering, Ocean University of China, 5 Yushan Road, Shinan District, Qingdao, Shandong Province 266003, China Department of Food Science, University of Massachusetts, Amherst, MA 01003, United States

a r t i c l e

i n f o

Article history: Received 22 July 2019 Received in revised form 2 August 2019 Accepted 7 August 2019 Available online 08 August 2019 Keywords: Carotenoid Delivery system Multilayer emulsions Stability Nutraceutical

a b s t r a c t Multilayer emulsions were formed by sequential electrostatic deposition of anionic (pectin) and cationic (chitosan) biopolymers onto anionic saponin-coated lipid droplets. These emulsions were then tested for their ability to encapsulate and protect a hydrophobic nutraceutical (astaxanthin). The impact of chitosan and pectin concentration, pH, and ionic strength on the formation and stability of the multilayer emulsions was examined. Multilayer emulsions containing small uniform particles were produced using 2.5% lipid droplets, 0.05% chitosan, and 0.0125% pectin. The physical stability of the astaxanthin-loaded emulsions after exposure to heating, pH, and NaCl was determined. The multilayer-coatings improved the chemical stability of the encapsulated astaxanthin, as well as the aggregation stability of the lipid droplets at elevated ionic strengths and temperatures. Astaxanthin degradation during storage was 3- to 4-fold slower in multilayer emulsions than conventional ones. The multilayer emulsions developed in this study may be useful for encapsulating, protecting, and delivering hydrophobic carotenoids, which may aid in the development of more efficacious functional foods, supplements, and medical foods. © 2019 Elsevier B.V. All rights reserved.

1. Introduction Astaxanthin (Ax), which is a member of the carotenoid family, is found in aquatic animals, such as shrimp, crabs, and salmon [1]. Previous studies have reported that astaxanthin has strong antioxidant properties, with singlet oxygen quenching activities N40- and 1000-fold higher than β-carotene and vitamin E, respectively [2]. In addition, it has been claimed that astaxanthin can prevent or treat certain diseases, including cataracts, age-related macular degeneration, inflammation, and cardiovascular disease [3]. Because of its highly unsaturated structure, astaxanthin has a tendency to chemically degrade when exposed to elevated temperatures, light, and oxidative conditions, which promotes color fading and loss of biological activity [4]. Moreover, the application of astaxanthin as a nutraceutical ingredient in functional foods and beverages is often limited due to its limited waterdispersibility. Finally, the relatively low oral bioavailability of carotenoids greatly limits their efficacy as nutraceuticals [5]. To improve the utilization of astaxanthin in functional foods, different strategies have been

⁎ Corresponding author. ⁎⁎ Correspondence to: D. Wang, College of Food Science and Engineering, Ocean University of China, 5 Yushan Road, Shinan District, Qingdao, Shandong Province 266003, China. E-mail addresses: [email protected] (D.J. McCleiments), [email protected] (D. Wang).

https://doi.org/10.1016/j.ijbiomac.2019.08.071 0141-8130/© 2019 Elsevier B.V. All rights reserved.

developed, including structural modification, microencapsulation, and food matrix design [6]. Emulsion-based delivery systems have been shown to be highly effective at improving the water-dispersibility, chemical stability, and oral bioavailability of carotenoids [7]. Emulsions are, however, unstable under many of the environments found in food and beverage products (such as certain pHs, ionic strengths, temperatures, and mechanical stresses), which can greatly reduce their applicability. Previous research has shown that the physicochemical stability of emulsions to environmental stresses can be improved by coating the lipid droplets with multiple layers of edible biopolymers [8]. The functionality of these “multilayer emulsions” can often be manipulated by controlling the number, type, and sequence of biopolymer layers used to coat the lipid droplets, as this allows the thickness, composition, charge, permeability, and integrity of the interfaces to be manipulated. The formation of multilayered biopolymer coatings around the lipid droplets in oil-inwFIJater emulsions therefore provides food technologists with a novel means of improving the stability and performance of many food and beverage products, as well to develop innovative colloidal delivery systems [8]. In this study, flaxseed oil was used as a carrier lipid for the carotenoids because it contains high levels of ω-3 fatty acids, which are known to provide health benefits [9]. Quillaja saponin was used as a natural small molecule surfactant to create the anionic lipid droplets that acted as templates for electrostatic deposition of the biopolymer

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coatings. This surfactant is extracted from the bark of the Quillaja saponaria Molina tree, and has been shown to be an effective emulsifier for oil-in-water emulsions [10,11]. Saponins consist of a hydrophobic triterpenoid or steroid backbone with one or more hydrophilic sugar chains attached [12]. These natural plant-based emulsifiers may therefore be suitable for replacing animal-based proteins and synthetic surfactants in food products [10,11,13]. The utilization of natural emulsifiers in the food industry is increasing due to consumer concerns about the environmental, ethical, and health issues associated with synthetic and animal-based alternatives [14,15]. In this study, two oppositely charged edible biopolymers, cationic chitosan and anionic pectin, were used to form multilayer coatings around anionic saponin-coated lipid droplets. Chitosan is a polysaccharide derived by alkaline N-deacetylation of chitin, which is a major component of crustacean shells such as crabs and shrimps [16,17]. Chitosan is widely used to create multilayer coatings because it is one of the few food-grade cationic polysaccharides available that can interact with anionic polysaccharides through electrostatic interactions. Pectin is a foodgrade anionic polysaccharide (pKa ≈ 3.5) usually isolated from plant cell walls, such as those in apples, citrus fruit, and beet pectin [18]. Pectin has been widely used as a building block for the fabrication of bioactive delivery systems with targeted and controlled-release properties [19]. The objective of the present study was to optimize the formation of stable multilayer-coated lipid droplets containing astaxanthin. The impact of the multilayer coatings on the physical and chemical stability of these emulsion-based delivery systems to environmental stresses commonly found in foods (such as pH changes, salt addition, and heating) was then evaluated. The results of this study may lead to the rational design and fabrication of more effective nutraceutical-loaded colloidal delivery systems for application in functional foods and beverages.

(2 min), which led to a clear solution. The oil and aqueous phases were then blended together for 2 min using a high-speed blender (M133/1281–0, Biospec. Product, Inc., ESGC, Switzerland) at 10,000 rpm. The resulting coarse emulsion was then passed threetimes through a microfluidizer at an operational pressure of 12,000 psi (Microfluidizer, M-110P, Microfluidics, Newton, MA USA). The resulting control emulsions and Ax-loaded emulsions were stored at 4 °C before being used. 2.3.2. Secondary emulsion The secondary emulsions (“Q-C bilayer emulsions”) were formed by adding 2.5 mL aliquots of primary emulsions to 2.5 mL aliquots of chitosan solutions (with varying initial concentrations). In addition, 5 mL aliquots of phosphate buffer solutions (pH 4.0) were added to the samples to dilute them. These samples were then vortexed for 1 min to ensure they were thoroughly mixed. 2.3.3. Tertiary emulsion The tertiary emulsions (“Q-C-P multilayer emulsions”) were formed by mixing secondary emulsions, phosphate buffer solutions (pH 4.0), and pectin solutions (varying initial concentrations, pH 4.0) at a volume ratio of 2:1:1. These samples were then vortexed for 1 min. 2.4. Particle size measurement

2. Materials and methods

The particle size distribution of the samples was measured using a static light scattering device (Mastersizer 2000, Malvern Instruments Ltd., Malvern, Worcestershire, UK). Initially, the samples were diluted with phosphate buffer solution (at the same pH as the sample) and then stirred to ensure they were homogeneous and to avoid multiple scattering effects. The refractive indices of the oil and aqueous phases used in the calculations were 1.479 and 1.333, respectively. The particle sizes are reported as volume-weighted mean diameters (d3,2) for all samples.

2.1. Materials

2.5. Particle charge measurements

Chitosan (Code 448,877, medium molecular weight (Mw = 1 to 20 kDa) with 75% deacetylation, viscosity (1% solution) 200–600 mPa) was obtained from Primex ehf (Reykjavik, Iceland. Pectin extracted from citrus peel was purchased from Sigma Chemical Company (St. Louis, MO, USA). Quillaja saponin (Q-Naturale 200®, 14% (w/w) active saponins) was kindly provided by Ingredion Inc. (Westchester, IL, USA). Flaxseed oil was provided by the Stepan Company (Bordentown, NJ, USA). Astaxanthin (N97%) was purchased from Sigma-Aldrich Co., Ltd. (St. Louis, MO, USA). Sodium azide was purchased from SigmaAldrich (St. Louis, MO). All organic solvents were of HPLC grade and other chemicals were of analytical grade.

Emulsions were diluted to a droplet concentration of approximately 0.005 wt% oil using 0.05 M phosphate buffer solution (at the same pH as the sample) prior to analysis. Diluted emulsions were then injected into the measurement chamber of an electrophoresis instruments. An electrophoresis instrument (Zetasizer Nano-ZS90, Malvern Instruments Ltd., Worcestershire, UK) was used to measure the surface-potential (zeta-potential) of the particles in the emulsions. The surface potential was determined by using a laser beam to measure the direction and velocity of particle movement in a well-defined electric field (150 V).

2.2. Stock solution preparation A 1 wt% chitosan solution was prepared by dispersing chitosan powder in 1% acetic acid and stirring overnight. A 0.5 wt% pectin solution was prepared by dissolving pectin powder in phosphate buffer (pH 4.0) and stirring overnight. When required, each of these stock biopolymer solutions was diluted to the required final concentration using buffer solution.

2.6. Turbidity measurements The turbidity of the biopolymer solutions was measured using an UV − visible spectrophotometer (Cary 100 UV–Vis, Agilent Technologies) with a cuvette (1 cm path length) at 600 nm. The pH of the chitosan and pectin solutions (0.1%, w/w) were adjusted by adding NaOH or HCl (1.0, 0.5, 0.1 M) to obtain values ranging from pH 2.0 to 7.0 and from 2.0 to 10.0, respectively. A 10 mM phosphate buffer solution was used as a blank [20]. 2.7. Confocal Laser Scanning Microscopy (CLSM) analysis

2.3. Sample preparation 2.3.1. Primary emulsion Initially, a stock emulsion was prepared by combining 10% wt% of oil phase (flaxseed oil with or without astaxanthin) with 90% wt% of aqueous phase (1% w/w Q-Naturale, 10 mM phosphate buffer, pH 7.0). The Ax-loaded oil phase was prepared by dispersing astaxanthin (0.5 mg/mL) in flaxseed oil by heating (50 °C, 2 h) and sonicating

The microstructures of the emulsions were characterized using a confocal fluorescence microscopy (Leica TCS SP5, Leica Microsystem, Mannheim, Germany) with a 10× eye piece and a 60× oil immersion objective lens operating in a fluorescence mode. Aliquots 15 μL Nile Red solution (1 mg/mL in ethanol) and 10 μL FITC (10 mg/mL in dimethyl sulfoxide) were mixed into 0.1 mL of the emulsion to dye the oil phase and proteins, respectively. The samples were excited at

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wavelengths of 543 and 605 nm and then intensity of the emitted fluorescence signal was measured at wavelengths of 488 and 515 nm for Nile red and FITC, respectively. The stained emulsion samples (4 μL) were transferred onto a microscope slide and then covered with a glass coverslip. The microstructural images were then captured and analyzed using the instrument software [21].

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times, and then centrifuged at 4000 rpm for 15 min. The absorbance of the collected dichloromethane layers was then determined using a UV–visible spectrophotometer at a wavelength of 474 nm (Cary 100 UV–Vis, Agilent Technologies). The fraction of astaxanthin remaining in the emulsions (C(t)/C(0)) was then measured, where C(0) and C(t) are the concentrations of Ax in the emulsion after 0 and t days storage, respectively.

2.8. Physical and chemical stability 2.9. Long storage stability 2.8.1. Thermal stability Emulsions were incubated in a preheated water bath at fixed temperatures (20, 30, 50, or 80 °C) for 30 min, then cooled to room temperature using ice bath, and then stored overnight at room temperature. 2.8.2. Salt stability Emulsions were stored for 12 h at 25 °C and then mixed with aqueous solutions containing different levels of NaCl to obtain a range of final salt concentrations (50, 100, 500, or 1000 mM). The samples were then vortexed for 1 min to ensure they were thoroughly mixed and then stored overnight [22]. 2.8.3. pH stability Emulsions were adjusted to a range of pH values (2 to 8) using HCl and/or NaOH solutions, and then stored overnight at ambient temperature before further analysis [23]. The physical stability of the emulsions was monitored by measuring changes in their particle size and overall appearance. 2.8.4. Color analysis Changes in the color of the Ax-loaded emulsions were monitored using an instrumental colorimeter to provide an indication of the chemical stability of the encapsulated astaxanthin (ColorFlex EZ, HunterLab, Reston, Virginia, USA). A fixed volume (10 mL) of emulsion was pipetted into a transparent disposable petri dish. The L* (lightness), a* (red to green), and b* (yellow to blue) measurements were recorded using a black cup as a background at room temperature. 2.8.5. Astaxanthin measurement The amount of astaxanthin present in the emulsions was also measured immediately after their preparation and after 15 days storage to provide information about the chemical stability of the carotenoid [24]. A fixed volume (2 mL) of Ax-loaded emulsion was mixed with 2 mL of dichloromethane and 2 mL of methanol, vortexed several

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2.10. Statistical analysis All data are expressed as mean and standard deviation of three measurements (n = 3). The differences between groups are assessed using a one-way ANOVA analysis with Tuckey's post hoc test (SPSS 20, Chicago, IL). A value of p b 0.05 was considered to be statistically significant. 3. Results and discussion 3.1. Influence of pH on electrical characteristics and aggregation state of biopolymers The surface potential and turbidity of the chitosan and pectin solutions were measured to provide information about the pHdependence of the electrical characteristics and aggregation state of the dissolved biopolymers. As shown in Fig. 1A, the chitosan molecules were positive for all pH values studied, with the magnitude of the surface potential depending on the pH of the surrounding solution. When the pH was increased from 2.0 to 5.0, the positive charge remained relatively high and constant (+25 mV), but it decreased appreciably when the pH was raised further. This effect is due to the fact that the amino groups on the chitosan have pKa values around 6.5, and therefore they lose their positive charge at higher pH values. Conversely, pectin molecules were negative across the entire pH range, but the magnitude of the

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charge was again pH-dependent. In particular, the surface potential remained relatively high and negative when the pH was reduced from pH 7.0 to 5.0, but then decreased appreciably when the pH was reduced further. This effect is due to the fact that the pKa value of the carboxylic acid groups on the pectin molecules is around 3.5 and so they become partially protonated at lower pH values [25]. The chitosan solutions remained optically clear from pH 2.0 to 6.0, but became strongly turbid at higher pH values, which can be attributed to aggregation of the chitosan molecules due to a reduction in the electrostatic repulsion between them (Fig. 1B). Conversely, there was a slight decrease in the turbidity of the pectin solutions when the pH was increased from 2.0 to 10.0, with the solutions going from cloudy (pH 2.0–3.0) to slightly turbid (pH 4.0–7.0) to clear (pH 8.0–10.0). These results suggest that the pectin molecules (or any insoluble pectin matter) tended to aggregate at lower pH values when they had little negative charge, but dissociate at higher pH values when they had a strong negative charge. 3.2. Influence of pH on the formation and properties of secondary emulsions The effect of pH on the particle size distributions of the saponincoated lipid droplets in the presence of chitosan is shown in Fig. 2A.

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With the exception of pH 2.0, all the emulsions contained a wide range of particle sizes suggesting that some aggregation of the individual lipid droplets had occurred. The largest particles were observed in the emulsions at pH 6.0 and 7.0, which may have been due to a reduction in the electrostatic repulsion between the chitosan molecules associated with their loss of positive charge at high pH values. Initially, the saponin-coated lipid droplets in the primary emulsions were negatively charged, which can be attributed to the anionic nature of the adsorbed saponin molecules. Under these circumstances, the cationic chitosan molecules in the surrounding solution can adsorb to the surfaces of the anionic lipid droplets through electrostatic attraction. At all pH values (Fig. 2C), the surface potential of the droplets in the secondary emulsions was positively charged, suggesting that the cationic chitosan molecules formed a polysaccharide coating around the anionic saponin-stabilized lipid droplets. Presumably, deprotonation of the amino groups on the chitosan molecules at high pH values reduced the magnitude of the positive charge on the outer chitosan layer [26]. As a result, the magnitude of the surface potential on the secondary emulsions decreased with increasing pH. Conversely, the surface potential of the chitosan-coated lipid droplets was sufficiently high (about +40 mV) to be stable at pH 4.0 and below. The chitosan-coated lipid droplets had almost no charge at pH 7.0 because the chitosan molecules

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lost their positive charge as the pH approached the pKa values of the amino groups [27]. Our results suggest that coating the lipid droplets with a layer of chitosan led to the formation of emulsions containing relatively small particles with high cationic charges (pH 4.0). 3.3. Influence of chitosan on the formation and properties of secondary emulsions The particle size distribution and mean particle diameter of the secondary emulsions was measured as a function of initial chitosan solution concentration (0–0.5 wt%) at pH 4.0. At initial chitosan solution concentrations from 0.01% to 0.08% (Fig. 3A), the secondary emulsions had monomodal distributions but they contained large particles (˃10 μm), which can be attributed to charge neutralization and bridging flocculation effects. At intermediate chitosan concentrations, the net droplet charge goes close to zero, which reduces the electrostatic repulsion between them. Moreover, the droplet surfaces are not fully covered with chitosan molecules, which means that a positive patch on the surface of one droplet can bind to a negative patch on the surface of another. When the initial chitosan

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solution concentration was increased further (0.1–0.5%) there was a decrease in the mean particle diameter, but the emulsions appeared to contain a broad range of particle sizes. This suggests that the droplets were fully coated by chitosan, which increased the steric and electrostatic repulsion between them, thereby reducing the degree of droplet flocculation. However, there still appeared to be some large flocs in the emulsions. Our results are consistent with those reported in previous studies on related systems [28,29]. The surface potential of the droplets in the emulsions went from strongly negative in the absence of chitosan to strongly positive in its presence. This effect can be attributed to the adsorption of cationic chitosan molecules onto the surfaces of the anionic saponin-coated lipid droplets due to a strong electrostatic attraction between them. A similar phenomenon has been reported by other authors for related emulsionbased systems [30]. The positive charge on the lipid droplets increases as the level of chitosan added increases because the cationic chitosan molecules progressively cover more and more of the anionic droplet surfaces. Eventually, the droplet surfaces are completely saturated by a layer of chitosan molecules and so the surface potential reaches a constant positive value.

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3.4. Influence of pectin on the formation and properties of tertiary emulsions After we had established the optimum chitosan concentration required to form secondary emulsions containing relatively small chitosan-coated lipid droplets, we examined the impact of adding anionic pectin to these systems to form tertiary emulsions containing pectin/chitosan-coated lipid droplets. The mean diameter and surface potential of the particles were measured after increasing amounts of pectin were added to the secondary emulsions (Fig. 4). The mean particle diameter remained relatively low from 0 to 0.05 wt% initial pectin, but then increased substantially as the pectin level was increased further, which is indicative of extensive droplet aggregation. A similar phenomenon has been reported by other researchers [31]. The surface potential of the particles in the emulsions was also measured as the pectin concentration was increased (Fig. 4C). The secondary emulsions, which contained chitosan-coated lipid droplets had a positive surface potential (+40 mV). As the amount of pectin added to these emulsions increased, the positive charge on the particles decreased, which can be attributed to the adsorption of anionic pectin molecules to the surfaces of the chitosan-coated lipid droplets. As a

result, there will have been a decrease in the electrostatic repulsion between the droplets, which could have led to some droplet aggregation. Moreover, there may have been some lipid droplets that were only partially covered by pectin molecules, leading to the presence of some anionic pectin patches and some cationic chitosan patches on their surfaces, which could promote bridging flocculation. It should be noted that full saturation of the droplet surfaces with pectin was not reached for the range of pectin concentrations used in this study. If saturation had occurred, then the droplets would have gained a strong net negative charge. Nevertheless, there are advantages to having colloidal delivery systems containing cationic particles. Previous work has reported that adsorption of cationic particles on target cell surfaces is rapid and independent of cell type [32]. Thus, the cationic particles in the multilayer emulsions developed in this study may be useful for delivering bioactive agents to anionic cell membranes. Based on our results, we used a final pectin concentration of 0.0125 wt% to prepare the tertiary emulsions in subsequent experiments since this led to relatively stable emulsions containing small cationic lipid droplets. These tertiary emulsions contained lipid droplets that were presumably coated by a multilayer consisting of anionic saponin, cationic chitosan, and some anionic pectin molecules.

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3.5. Microstructures of multilayer emulsions The microstructures of the Ax-loaded emulsions (pH 4) were characterized by confocal fluorescence microscopy (Fig. 5). The lipid droplets were stained red and the proteins were stained green to differentiate them. The microscopy images showed that the primary emulsions contained small lipid droplets that were fairly evenly spread throughout the sample. There appeared to be a slight increase in particle size for the secondary emulsions and a more pronounced increase for the tertiary emulsions. These results suggest that a limited amount of droplet aggregation occurred during formation of the secondary and tertiary emulsions. These results agree with the particle size data obtained by laser diffraction (see previous section), which also showed

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that the particle size decreased in the following order: tertiary N secondary N primary emulsions. Visual observation of the emulsions after storage indicated that the primary and secondary emulsions had fairly good stability to creaming, whereas the tertiary emulsions exhibited some phase separation. This result again supports the fact that there was some droplet aggregation in the multilayer emulsions. 3.6. Physical and chemical stability 3.6.1. Thermal stability The effect of thermal treatments (30, 50, and 80 °C for 30 min) on the particle size, surface potential, color, and appearance of the three emulsions were measured (Fig. 6). Little change was observed in the size of

Fig. 5. Confocal laser scanning micrographs of the Ax-loaded primary emulsions (A1-A3), secondary emulsions coated with chitosan (0.05%) (B1-B3), and tertiary emulsions coated with pectin (0.0125%) (C1-C3). The subscripts refer to: (1) lipid stain; (2) protein stain; (3) both lipid and protein stain.

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aggregation [33]. In addition, heating the emulsions may have promoted conformational changes in the adsorbed biopolymers that promoted their interfacial rearrangement. The tertiary emulsions exhibited a similar particle size-temperature dependence as the secondary emulsions: there was little change from 20 to 50 °C but then a steep increase as the temperature was raised higher. This might be expected because the level of pectin present in the outer interfacial layer would be expected to be quite small. Overall,

the particles within the primary emulsions after storage at elevated temperatures, indicating that they had good thermal-stability. There was, however, a substantial increase in the mean particle diameter in the secondary emulsion with rising temperature. For instance, the mean particle diameter (d32) increased from around 0.48 μm at 20 °C to 0.68 μm at 50 °C and then to 0.90 μm at 80 °C. This phenomenon may have occurred because raising the temperature enhanced the collision frequency of the lipid droplets, thereby promoting their

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these results suggest that there was some change in the composition or structural organization of the biopolymer layers around the lipid droplets at higher temperatures, which promoted droplet aggregation. Similar results have been reported for other multilayer emulsions [22]. There was, however, no significant (p N 0.05) impact of temperature on the surface potential (Fig. 6B) of the lipid droplets in any of the samples, which suggests that there were only minor changes in the interfacial composition upon heating.

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Visible observations of the emulsions indicated that there were no changes in the appearances of the secondary and tertiary emulsions after heating. Conversely, considerable oil leakage was observed in the primary emulsions, as seen by the presence of a liquid oil layer on the surfaces of the emulsions after heating. These results suggest that using saponins alone is insufficient to give good thermal stability to the emulsions, but coating the lipid droplets with an additional one or two layers of biopolymers greatly improves their thermal stability. In a

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related study, oil droplets coated by hydrolyzed rice glutelin and quillaja saponin were found to be more resistant to thermal processing than oil droplets coated by quillaja saponin alone [21], which was attributed to strong steric and electrostatic repulsion due to the biopolymer layer. Changes in the color of the Ax-loaded emulsions were monitored when they were held at a range of incubation temperatures (Fig. 6). The Ax-loaded emulsions had a uniform pink appearance. There were slight changes in the lightness (L*) of all the emulsions during storage.

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In particular, the redness (a*) value of the emulsions decreased moderately as the storage temperature was raised, which is indicative of some astaxanthin degradation. The secondary and tertiary emulsions also exhibited some loss of redness during incubation, suggesting that neither coating could completely inhibit heat-induced carotenoid degradation. Overall, our results suggest that coating the lipid droplets with chitosan is not particularly effective at protecting the astaxanthin from thermal degradation.

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3.6.2. Salt stability In this series of experiments, the impact of adding salt (NaCl) on the physicochemical stability of the emulsions was investigated. For the primary emulsions, the mean particle diameter increased slightly when the salt concentration was raised from 0 to 50 mM, but then increased substantially when the salt level was raised further, indicating that extensive droplet aggregation occurred at high ionic strengths. For the secondary emulsions, the mean particle diameter remained relatively constant from 0 to 100 mM salt, but then increased appreciably when the salt level was raised further. This suggests that the secondary emulsion was somewhat more stable than the primary emulsion to salt addition, but that aggregation still occurred at sufficiently high salt levels. For the tertiary emulsions, the mean particle diameter remained relatively low at all salt levels employed, indicating that they were highly resistant to salt addition. The good salt-stability of these emulsions was probably a result of the thick and highly charged biopolymer coating surrounding the lipid droplets, which increased the steric and electrostatic repulsion. The surface potential of the particles in the primary, secondary, and tertiary emulsions did not change appreciably with salt addition (Fig. 7B), which suggests that the interfacial composition did not change appreciably. Instrumental colorimetry measurements were used to quantify the impact of salt addition on the appearance of the three types of emulsion. There was a substantial decrease in the lightness (L*) and alteration in the color coordinates (a* and b*) of the primary emulsion when NaCl was added, which may have been due to changes in the light scattering efficiency of the lipid droplets when they aggregated in the presence of salt. Conversely, there was little change in the tristimulus color coordinates of the secondary and tertiary emulsions after salt addition, suggesting they were more stable to salt ions. Our results are in good agreement with previous studies on related systems [25].

3.6.3. pH stability In this section, the impact of pH on the physicochemical stability of the three emulsions was determined (Fig. 8). For the primary emulsions, the particles were relatively small from pH 8.0 to 3.0, but increased substantially when the pH was reduced to 2.0 (Fig. 8A). This effect can be attributed to the reduction in the magnitude of the negative charge on the saponin molecules adsorbed to the lipid droplet surfaces as the pH was reduced (Fig. 8B). For the secondary emulsions, the particle size was relatively small at pH 4.0 and 5.0, but increased appreciably when the pH was raised or lowered (Fig. 8A). The instability of the emulsions at relatively high pH values can be attributed to the loss of the positive charge on the outer chitosan layer (Fig. 8B). Conversely, the instability observed at low pH values may have been because the saponin molecules lost some of their negative charge, and so the chitosan molecules could not stick to their surfaces as strongly. For the tertiary emulsions, the particle size remained relatively small and constant from pH 3.0 to 5.0, but increased appreciably at higher or lower pH values. Again, this effect may have been due to a breakdown of the interfacial coatings under highly acidic or basic conditions. Instrumental colorimetry measurements were used to assess the impact of pH on the appearance of the three emulsions (Figs. 7C-E). For the primary emulsions, the lightness remained relatively constant from pH 8.0 to 3.0, but decreased appreciably at pH 2.0, which may have been due to a change in the light scattering efficiency of the droplets when they aggregated under acidic conditions. For both multilayer emulsions, there was a slight decrease in lightness, but little change in a* and b* values, when the pH was increased from 2.0 to 8.0. Overall, these results suggest that coating the lipid droplets with biopolymers improved their stability over some pH ranges but not others.

3.7. Physical and chemical stability during long storage Astaxanthin is a carotenoid with a highly unsaturated structure that is highly sensitive to oxidation. For this reason, we examined the impact of environmental stresses on the physical and chemical stability of the emulsions during 15 days storage at 37 °C (Fig. 9). The primary emulsion was highly stable to droplet aggregation during storage, i.e., the mean particle diameter remained relatively constant (Fig. 9A). Conversely, there was a moderate but steady increase in the mean particle diameter in the secondary and tertiary emulsions during storage. Even so, this increase was not large enough to lead to visible phase separation. The surface potential of the particles in the primary emulsions remained relatively high and negative during the first 9 days of storage, but then the magnitude of the negative charge decreased appreciably, suggesting there was some change in the interfacial composition of this system. For the secondary and tertiary emulsions, the surface potential remained relatively constant throughout storage, indicating that the surface composition and/or structure remained relatively constant. The chemical stability of the astaxanthin was also measured to determine the extent of carotenoid degradation during storage (Fig. 9CF). Instrumental colorimetry showed that the a* and b* values of the primary emulsions decreased rapidly during the first few days of storage, whereas those of the secondary and tertiary emulsions only decreased quite slowly, which suggested that the multilayer coatings protected the encapsulated carotenoids from degradation. Measurements of the fraction of astaxanthin (C/C0) remaining in the emulsions during storage showed that there its degradation was much faster in the primary emulsions than in the secondary or tertiary emulsions. Previous studies have also reported that a cationic outer chitosan layer can inhibit the oxidation of emulsified oils, which was attributed to their ability to repel pro-oxidative metal cations (Klinkesorn, Sophanodora, Chinachoti, McClements, & Decker, 2005; Ogawa, Decker, & McClements, 2003). The impact of storage on the overall appearance of the three emulsions was also measured (Fig. 9G). After 15 days storage at 37 °C, all of the emulsions had fairly similar appearances, with none of the emulsions exhibiting any obvious phase separation. However, the multilayer emulsions did appear to be more yellow in color than the primary emulsions. 4. Conclusion All-natural emulsion-based delivery systems have been created for hydrophobic carotenoids by coating anionic saponin-stabilized lipid droplets with cationic chitosan then anionic pectin. The optimum conditions for producing multilayer emulsions containing small highly charged droplets were established for the two biopolymers used. The multilayer emulsions had better stability than conventional single layer emulsions under certain pH, salt, and temperature conditions. Moreover, the multilayer biopolymer coatings protected the carotenoids from chemical degradation during storage, which may be advantageous for extending their shelf-life and efficacy. In summary, this research provides valuable new information that should aid the development of biopolymer-coated lipid droplets that can encapsulate, protect, and deliver labile hydrophobic nutraceuticals. These delivery systems may be useful for application in commercial food, supplement, and pharmaceutical products. Notes The authors declare no competing financial interest.

Fig. 9. Changes in mean particle diameter (A), surface potential (B), color coordinates (C-E), Ax degradation (F), and appearance (G) of Ax-loaded primary, secondary (0.05% chitosan), and tertiary (0.0125% pectin) emulsions during storage at 37 °C.

C. Liu et al. / International Journal of Biological Macromolecules 140 (2019) 985–997

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