Article pubs.acs.org/Langmuir
Oil-in-Water Emulsion System Stabilized by Protein-Coated Nanoemulsion Droplets Aiqian Ye,* Xangqian Zhu, and Harjinder Singh* Riddet Institute, Massey University, Private Bag 11222, Palmerston North 4442, New Zealand
ABSTRACT: Nanoemulsion droplets (average size of about 150 nm) coated with micellar casein were used as an emulsifying agent to stabilize oil (n-hexadecane)-in-water emulsions. We found that these nanodroplets adsorbed at the oil−water interface and formed stable emulsions, with the size of the droplet-stabilized emulsions being dependent on the concentration of nanodroplets in the dispersions. Stable emulsions were still formed at low concentrations, even though the interface was not fully covered by the nanodroplets. The nanodroplets fully covered the interface at higher concentrations, resulting in a transition from a thick interfacial layer to a multilayer containing a network of assembled nanodroplets. Because of their soft and elastic nature, the adsorbed nanodroplets showed strong deformation at the oil−water interface. The morphology of the interfacial nanodroplets was dependent on their location inside the network interfacial layer. The adsorption and deformation of the droplets at the oil−water interface and the formation of network structures, as observed in the present study, provide new, useful fundamental knowledge with potential applications for microencapsulation and emulsion stabilization.
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INTRODUCTION The adsorption of solid particles at air−water and oil−water interfaces has attracted considerable research attention over the past decade, largely because of the many potential applications in the stabilization of foams and emulsions. The assembly of adsorbed colloidal particles at the surface is expected to provide long-term stability.1,2 Most of the particles selected to produce particle-stabilized emulsions have been synthetic, e.g., polymer latices, silica, metal oxides, and polymeric microgel particles.3−6 Recently, nonsurface-activated CaCO3 nanoparticles have been used to stabilize oil-in-water emulsions through in situ interaction with the anionic surfactant sodium dodecyl sulfate (SDS).7 Moreover, stimuli-sensitive soft microgel particles have been employed as stabilizers for emulsions. These microgel particles are able to adsorb at the oil−water interface and show considerably different behavior from the commonly used Pickering stabilizers, which are solid particles.2,8 Because they are soft and deformable, they present some advantages as emulsifying agents; for example, the deformability of the soft particles at the interface makes it possible to cover a larger interfacial area and form a densely packed compliant shell, which is important for some applications, e.g., in fabricating capsules and preventing coalescence on severe droplet deformation.6,9,10 However, only a few natural materials have been reported to act as particle stabilizers.11−13 Relatively less published research is directly applicable to foods because many of these particles are not permissible in foods. Murray et al.13 © 2013 American Chemical Society
reported on the adsorption behavior of mixtures of surfaceactive polymers and surface-active food-grade particles, such as oil droplets, cellulose, and starch particulates. Recently, some other particles derived from food materials, e.g., starch nanocrystals, flavonoids, nanoparticles, nanoparticle−protein complexes, and lactoferrin nanoparticles, were applied in the formation of the model emulsions.14−19 Proteins are the most important emulsifying agents for making edible oil-in-water emulsions, because of their amphiphilic character and high surface activity. They impart emulsion stability by a combination of electrostatic and steric mechanisms.20,21 One such protein-rich emulsifying agent is the casein micelle that is present in milk.22,23 Casein micelles are considered to be colloidal particles because they consist of aggregated casein proteins with a stabilizing surface layer. κCasein is predominantly on the surface of the casein micelle, βcasein is mostly present in the interior, and αs1- and αs2-caseins are found throughout the micelle structure. It has been reported that the available κ-casein covers about one-third of the surface of the casein micelles and is heterogeneously distributed on the surface, and that the other caseins may share the surface with κ-casein. Casein micelles adsorb at the oil− water interface and then spread along the interface.24,25 The Received: September 9, 2013 Revised: October 29, 2013 Published: October 31, 2013 14403
dx.doi.org/10.1021/la403493y | Langmuir 2013, 29, 14403−14410
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Article
(Nanosizer ZS, Malvern Instruments Ltd., Malvern, Worcestershire, U.K.)), which was similar to the size of the casein micelles in milk.27 However, the average particle size (d32) of the casein micelles is ∼110 nm when the size is determined by static light scattering (Malvern MasterSizer 2000, Malvern Instruments Ltd., Malvern, Worcestershire, U.K.). This indicated that intact casein micelles remained in the dispersion. n-Hexadecane was purchased from Sigma Chemical Co. (St. Louis, MO). All the chemicals used were of analytical grade and were obtained from either BDH Chemicals (BDH Ltd., Poole, U.K.) or Sigma−Aldrich (St. Louis, MO), unless specified otherwise. Preparation of Emulsions. Casein micelle dispersions were prepared by adding micellar casein powder to 10 mM pH 7 phosphate buffer and then stirring for 60 min at room temperature to ensure complete dispersion. The dispersions were adjusted to pH 7.0. Appropriate quantities of n-hexadecane were then mixed with the protein dispersions to give 20% n-hexadecane in the mixtures, which were then homogenized in a microfluidizer (M-110EH; Microfluidics Corporation, Newton, MA) at a pressure of 300 MPa to form the nanoemulsions. Appropriate quantities of n-hexadecane were mixed with the nanoemulsions to give 20% n-hexadecane and 2−16% nanoemulsion in the final emulsions. The mixtures of n-hexadecane and nanoemulsion were then homogenized in a rotor−stator Ultra Turrax homogenizer (10 000 rev/min for 2 min) (Diax 600; Heidolph, Germany) to form the final emulsions. Characterization. A Malvern MasterSizer 2000 (Malvern Instruments Ltd., Malvern, Worcestershire, UK) was used to determine the average diameter (d32) of the emulsion droplets. The relative refractive index (N), i.e., the ratio of the refractive index of the emulsion droplets (1.456) to that of the dispersion medium (1.33), was 1.095. The absorbance value of the emulsion droplets was 0.001. Droplet size measurements are reported as average diameters, d32. The d32 is defined as ∑nidi3/∑nidi2, where ni is the number of particles with diameter di. Mean particle diameters were calculated as the average of duplicate measurements. The protein concentrations of the aqueous phase of the emulsions were characterized as that the emulsions were centrifuged at 45000g for 30 min at 20 °C, the subnatants were then filtered through 0.22 μm filters (Millipore). The protein concentrations of emulsions and the filtrate were determined by the Kjedahl method (1026 Distilling Unit and 1007 Digestor Blorck, Tecator AB, Hoganas, Sweden). The microstructure of the samples was examined using a confocal laser scanning microscopy (CLSM) system (Leica DM6000B, Heidelberg, Germany) with a 63 mm oil immersion objective lens. Nile Blue was used to stain for oil (argon laser with an excitation line of 488 nm) whereas Fast Green was used to stain for protein in the bolus (He−Ne laser with excitation at 633 nm). Samples (1 mL) were stained with 1.0% (wt/vol) Nile Blue and 1.0% (wt/vol) Fast Green (fluorescent dye), placed on a concave confocal microscope slide (Sail; Sailing Medical-Lab Industries Co. Ltd., Suzhou, China), covered with a coverslip, and finally examined with a 63× magnification lens using the microscope. The microstructure of the samples was also studied using transmission electron microscopy (TEM). A sample was mixed with warm melted (35−40 °C) 3% low-temperature-gelling agarose in a 1:1 ratio. This solution was poured on to a microscope slide, allowed to set, and chopped into 1 mm3 cubes. The cubes were put into a bijoux bottle containing 3% glutaraldehyde in 0.2 M sodium cacodylate buffer. This was kept at 5 °C for 24 h. The samples with glutaraldehyde were then rinsed twice with 0.2 M sodium cacodylate buffer over 2 h. The agarose-embedded samples were placed in 1% osmium tetroxide (1 mL) overnight at room temperature. The samples were rinsed twice with distilled water, placed in 1% uranyl acetate (1 mL) for 30 min, and then rinsed twice with distilled water. The dehydration process was carried out at 5 °C in 25% acetone (15 min) and then in 50, 70, and 90% acetone (for 30 min each), followed by 100% acetone (three changes over 90 min). The acetone was replaced with Procure 812 embedding resin, and the samples were put on rollers for 24 h. Sample cubes were placed into an embedding capsule and were cured at 60 °C for 48 h. The embedded samples
redistribution and the spreading of casein micelles at the interface are dependent on the conditions of adsorption, e.g., homogenization pressure, protein concentration, and other emulsifying agents in the system. Murray et al.13 used emulsion droplets coated with protein to strengthen the adsorbed film and to provide enhanced stability for air bubbles, although the emulsion droplets may not have been completely solid. It was considered that the droplet particles had the advantage that they were relatively welldefined in terms of their size, shape, and surface properties. However, the emulsion droplets did not act as a foaming agent because the air bubbles were stabilized by the protein film at the air−water interface. The emulsion droplets associated with the adsorbed protein film under certain conditions, such as at low pH. This association led to the formation of a strong adsorbed film to promote foam stability. In some cases, proteincoated emulsion droplets can become solid particles, e.g., when solid fat such as milk fat or palm oil is used, depending on the environmental temperature. Recently, the use of proteinstabilized emulsion droplets as emulsifying agents to stabilize oil-in-water emulsions was demonstrated in a patent application.26 In this work, we developed a novel stable emulsion system that was stabilized by nanoscale emulsion droplets that were coated with colloidal protein particles (casein micelles). The adsorption and surface properties of both the casein micelles and the casein-micelle-coated emulsion droplets were examined. The arrangement of the casein micelles and caseinmicelle-coated droplets at n-hexadecane droplet interfaces was probed via microscopy. The influence of the particle concentration and the oil/water ratio is discussed.
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EXPERIMENTAL SECTION
Materials. Micellar casein powder was obtained from the American Casein Company (Burlington, NJ). Its composition was as follows: protein 85%, fat 2.1%, ash 9.5%, lactose 3.2% on a dry solids basis, and moisture 4.8%. It was characterized using sodium dodecyl sulfatepolyacrylamide gel electrophoresis (SDS-PAGE) (Figure 1), which showed that only caseins were present. The average particle size of the micellar casein dispersed in 10 mM pH 7 phosphate buffer was found to be ∼200 nm (determined using dynamic light scattering,
Figure 1. SDS-PAGE pattern of micellar casein dispersion. 14404
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were then sectioned to a thickness of 90 nm using a Reichert Ultracut microtome. These sections were mounted on 3 mm copper grids and stained with lead citrate before examination in a Philips transmission electron microscope (NL-5600 MD; Philips, Eindhoven, The Netherlands) at an acceleration voltage of 60 kV. The ζ-potential values of the emulsion samples were measured using a laser Doppler velocimetry and phase analysis light scattering (M3-PALS) technique and a Malvern Zetasizer Nano ZS instrument (Malvern Instruments Ltd.). The samples were diluted to 100 times with Milli-Q water, placed in the electrophoretic mobility cell, and analyzed at a scattering angle of 173° using the Malvern Zetasizer Nano ZS. The effective electric field, E, applied in the measurement cell was between 50 and 150 V depending on the ionic strength of the samples. The overall electrophoretic mobility, μ, was calculated at 20 °C according to eq 1 (for a spherical particle). Y = μω /E
(1) −1
where Y is the particle velocity (m/s) and ω is the frequency (s ) of the applied electric field. The ζ-potential (mV) was then calculated using the Smoluchowski equation: μ = ζε/η
(2)
where ε is the electric permittivity of the solvent (F/m) and η is the solvent viscosity (Pa·s). An individual ζ-potential measurement was calculated from the mean of two runs and the standard deviation of at least 10 readings from an individual sample. A contact angle and surface tension meter (KSV Instruments Ltd., Helsinki, Finland) was used to examine the time evolution of the interfacial tension of protein and protein-coated emulsion droplets via the pendant drop method. The tensiometer operated in a volumecontrolled regime, via continuous measurement of the drop area and volume, approximated by a Laplacian profile. The protein solution or protein-stabilized emulsion was the drop phase against pure nhexadecane. The volume of the drop was about 8 μL. The shape of a drop hanging from a syringe tip was determined from the balance of forces, which included the surface tension of the liquid. The interfacial tension at the liquid interface can be related to the drop shape through eq 3: Δρ × g × R 0 γ= β
Figure 2. (a) Average droplet size (d32) (■) and protein concentration in the aqueous phase (□) of the nanoemulsions formed using micellar casein; (b) droplet size distribution of the nanoemulsion formed at a protein concentration of 40 mg/mL.
(3)
where γ is the interfacial tension, Δρ is the difference in density between fluids at the interface, g is the gravitational constant, R0 is the radius of drop curvature at the apex, and β is the shape factor. The shape factor can be defined through the Young−Laplace equation.
because of flocculation, with the casein micelles bridging between the droplets.28 At higher concentrations, the size of the emulsion droplets remained constant because the excess casein micelles stayed in the aqueous phase. Confocal microscopy of the emulsion formed at a casein micelle concentration of 40 mg/mL indicated a mostly uniform size distribution of small droplets; some droplets appeared to be flocculated (Figure 3). TEM showed clearly that almost all the casein micelles adsorbed on the surface of the emulsion droplets, which had fairly even size distributions (Figure 4). Most of the micelles present at the droplet surface were not intact and appeared to be uneven and spread around the droplet surface (Figure 4c). It is likely that, after adsorption, the casein micelle dissociated and spread along the interface, adjusting its structure to cover the maximum surface area. This resulted in the casein micelles forming a dense and continuous interfacial layer around the emulsion droplets at a relatively low protein concentration. Dissociation and spreading of casein micelles on the surface of fat globules have been observed in previous studies on recombined milk and homogenized milk.25,29 The spreading may have been driven by the thermodynamically favorable changes in the interfacial free energy, which appears to be sufficient to disrupt micelles.30 Another possibility is that homogenization caused disruption of
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RESULTS AND DISCUSSION Formation of Casein-Micelle-Stabilized Nanoemulsion Droplets. Figure 2a shows the diameters of n-hexadecane emulsion droplets stabilized with casein micelles as a function of the total and aqueous phase protein concentrations. The droplet size decreased with increasing protein (mainly casein micelle) concentration in the emulsion system. The average droplet size was about 150 nm when the protein concentration was 40 mg/mL and remained constant above this concentration, indicating the formation of a stable nanoemulsion at a protein concentration of 40 mg/mL; this emulsion showed a narrow droplet size distribution with sizes in the range 50−800 nm (Figure 2b). Very small amounts of protein remained in the aqueous phase (6%, Figure 6), some of the nonadsorbed droplets may interact with adsorbed droplets, resulting in flocs, which may form a dense multilayer with a three-dimensional network at the interface. This multilayer can provide additional stabilization and may play an important role in the stability of particle-stabilized emulsions.1 In a Pickering emulsion, solid particles adsorb and form a thick surface layer, in which the structure of the surface layer is dependent mainly on the particle size and the hydrophobicity, in that the particles need to be partly wetted by both phases.36 Significant deformation of the nanodroplets along the interface of the large droplets, observed in the present study, suggests a different adsorption behavior of soft particles from that of the solid, hard particles used in Pickering emulsion systems, in which the deformation of the solid particles at the interface is negligible. However, the deformation or partial deformation of the particles at the interface has been observed recently for soft particles, such as cross-linked polymeric hydromicrogels formed by NiPAm and BIS, the ligand shell of gold nanoparticles, and copolymer-capped magnetite nanoparticles.6,9,10 Geisel et al.9 intuitively described the deformation of microgel particles at the interface. The equilibrium shape of the microgel particles is given by the balance among the solvations of the hydrogel in the oil and water phases, their interfacial activity, and the internal elasticity of the particles. The particle is stretched radially until the energy gain from the interfacial activity is balanced by the internal elastic deformation of the hydrogel. Compared with these deformable microgel particles and other soft particles, the n-hexadecane nanodroplets used in our study would be softer and more elastic, which could result in more pronounced deformation at the interface, as displayed in Figure 9. The deformation of there nanodroplets is probably driven by a tendency to maximize the interaction of the casein aggregates (adsorbed at nanodroplet surface) with the oil/water interface via the bridging of protein between pairs of interfaces (nanodroplet/droplet). It can be estimated that the extent of deformation is dependent on the relative ratio of the droplet sizes, i.e., nanodroplet:large droplet, the interfacial tension, and the hardness of the interfacial nanodroplets. Furthermore, the deformation may be influenced by the microstructure of the surface layer or the presence of other droplets within the surface layer if a multilayer or three-dimensional network is formed.
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AUTHOR INFORMATION
Corresponding Authors
*A. Ye. E-mail: [email protected]. Tel: 64-6-3505072. *H. Singh. E-mail: [email protected]. Tel: 64-6-3504401. Notes
The authors declare no competing financial interest.
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REFERENCES
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(6) Schmidt, S.; Liu, T.; Rütten, S.; Phan, K.-H.; Möller, M.; Richtering, W. Influence of Microgel Architecture and Oil Polarity on Stabilization of Emulsions by Stimuli-Sensitive Core-Shell Poly(Nisopropylacrylamide-co-methacrylic acid) Microgels: Mickering versus Pickering Behavior? Langmuir 2011, 27, 9801−9806. (7) Cui, Z. G.; Cui, Y. Z.; Cui, C. F.; Chen, Z.; Binks, B. P. Aqueous Foams Stabilized by in situ Surface Activation of CaCO3 Nanoparticles via Adsorption of Anionic Surfactant. Langmuir 2010, 26, 12567− 12574. (8) Brugger, B.; Rosen, B. A.; Richtering, W. Microgels as StimuliResponsive Stabilizers for Emulsions. Langmuir 2008, 24, 12202− 12208. (9) Geisel, K.; Isa, L.; Richtering, W. Unraveling the 3D Localization and Deformation of Responsive Microgels at Oil/Water Interfaces: A Step Forward in Understanding Soft Emulsion Stabilizers. Langmuir 2012, 28, 15770−15776. (10) Richtering, W. Responsive Emulsions Stabilized by StimuliSensitive Microgels: Emulsions with Special Non-Pickering Properties. Langmuir 2013, 28, 17218−17229. (11) Russell, T. J.; Lin, Y.; Boker, A.; Su, L.; Carl, P.; Zettl, H.; He, J. B.; Sill, K.; Tangirala, R.; Emrick, T.; Littrell, K.; Thiyagarajan, P.; Cookson, D.; Fery, A.; Wang, Q.; Russell, T. P. Self-Assembly and Cross-Linking of Bionanoparticles at Liquid-Liquid Interfaces. Angew. Chem., Int. Ed. 2005, 44, 2420−2426. (12) Binks, B. P.; Clint, J. H.; Mackenzie, G.; Simcock, C.; Whitby, C. P. Naturally Occurring Spore Particles at Planar Fluid Interfaces and in Emulsions. Langmuir 2005, 21, 8161−8167. (13) Murray, B. S.; Durga, K.; Yusoff, A.; Stoyanov, S. D. Stabilization of Foams and Emulsions by Mixtures of Surface Active Food-Grade Particles and Proteins. Food Hydrocolloids 2011, 25, 627−638. (14) Li, C.; Sun, P. D.; Yang, C. Emulsion Stabilized by Starch Nanocrystals. Starch/Starke 2012, 64, 497−502. (15) Luo, Z. J.; Murray, B. S.; Ross, A. L.; Povey, M. J. W.; Morgan, M. R. A.; Day, A. J. Particle-Stabilizing Effects of Flavonoids at the OilWater Interface. J. Agric. Food Chem. 2011, 59, 2636−2645. (16) Rana, S.; Yu, X.; Patra, D.; Moyano, D. F.; Miranda, O. R.; Hussain, I.; Rotello, V. M. Control of Surface Tension at Liquid-Liquid Interfaces Using Nanoparticles and Nanoparticle-Protein Complexes. Langmuir 2012, 28, 2023−2027. (17) Rayner, M.; Timgren, A.; Sjoo, M.; Dejmek, P. Quinoa Starch Granules: A Candidate for Stabilising Food-Grade Pickering Emulsions. J. Sci. Food Agri. 2012, 92, 1841−1847. (18) Shimoni, G.; Levi, C. S.; Tal, S. L.; Lesmes, U. Emulsions Stabilization by Lactoferrin Nano-Particles under in Vitro Digestion Conditions. Food Hydrocolloids 2013, 33, 264−272. (19) Tzoumaki, M. V.; Moschakis, T.; Kiosseoglou, V.; Biliaderis, C. G. Oil-in-Water Emulsions Stabilized by Chitin Nanocrystal Particles. Food Hydrocolloids 2011, 25, 1521−1529. (20) Dickinson, E. Proteins at Interfaces and in Emulsions Stability, Rheology and Interactions. J. Chem. Soc., Faraday Trans. 1998, 94, 1657−1669. (21) McClements, D. J. Food Emulsions: Principles, Practices, and Techniques, 2nd ed.; CRC Press: Boca Raton, FL, 2005. (22) Dalgleish, D. G.; Corredig, M. The Structure of the Casein Micelle of Milk and Its Changes During Processing. Annu. Rev. Food Sci. Technol. 2012, 3, 449−467. (23) Singh, H. Aspects of Milk-Protein-Stabilised Emulsions. Food Hydrocolloids 2011, 25, 1938−1944. (24) Walstra, P. On the Stability of Casein Micelles. J. Dairy Sci. 1990, 73, 1965−1979. (25) Sharma, R.; Singh, H.; Taylor, M. W. Composition and Structure of Fat Globule Surface Layers in Recombined Milk. J. Food Sci. 1996, 61, 28−32. (26) Singh, H.; Ye, A.; Zhu, X. Emulsion. PCT Patent Application WO2011/136662 A1, April 26, 2011. (27) Anema, S. G.; Klostermeyer, H. Heat-Induced, pH-Dependent Dissociation of Casein Micelles on Heating Reconstituted Skim Milk at Temperatures Below 100 Degrees C. J. Agric. Food Chem. 1997, 45, 1108−1115.
(28) Ye, A. Functional Properties of Milk Protein Concentrates: Emulsifying Properties, Adsorption and Stability of Emulsions. Int. Dairy J. 2011, 21, 14−20. (29) Ye, A.; Singh, H.; Anema, S. Changes in the Surface Protein of the Fat Globules during Homogenization and Heat Treatment of Concentrated Milk. J. Dairy Res. 2008, 75, 347−353. (30) Walstra, P.; Oortwijn, H. The Membranes of Recombined Fat Globules. III. Mode of Formation. Neth. Milk Dairy J. 1982, 36, 103− 113. (31) Walstra, P. On the Stability of Casein Micelles. J. Dairy Sci. 1990, 73, 1965−1979. (32) Dalgleish, D. G. On the Structural Models of Bovine Casein Micelles − Review and Possible Improvements. Soft Matter 2011, 7, 2265−2272. (33) Dickinson, E. Food Emulsions and Foams: Stabilization by Particles. Curr. Opin. Colloid Interface Sci. 2010, 15, 40−49. (34) Hunter, T. N.; Pugh, R. J.; Franks, G. V.; Jameson, G. J. The Role of Particles in Stabilising Foams and Emulsions. Adv. Colloid Interface Sci. 2008, 123, 57−81. (35) Horozov, T. S.; Binks, B. P. Particle-Stabilized Emulsions: A Bilayer or a Bridging Monolayer? Angew. Chem., Int. Ed. 2006, 45, 773−776. (36) Leal-Calderon, F.; Schmitt, V. Solid-Stabilized Emulsions. Curr. Opin. Colloid Interface Sci. 2008, 13, 217−227.
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Oil-in-Water Emulsion System Stabilized by Protein-Coated Nanoemulsion Droplets Aiqian Ye,* Xangqian Zhu, and Harjinder Singh* Riddet Institute, Massey University, Private Bag 11222, Palmerston North 4442, New Zealand
ABSTRACT: Nanoemulsion droplets (average size of about 150 nm) coated with micellar casein were used as an emulsifying agent to stabilize oil (n-hexadecane)-in-water emulsions. We found that these nanodroplets adsorbed at the oil−water interface and formed stable emulsions, with the size of the droplet-stabilized emulsions being dependent on the concentration of nanodroplets in the dispersions. Stable emulsions were still formed at low concentrations, even though the interface was not fully covered by the nanodroplets. The nanodroplets fully covered the interface at higher concentrations, resulting in a transition from a thick interfacial layer to a multilayer containing a network of assembled nanodroplets. Because of their soft and elastic nature, the adsorbed nanodroplets showed strong deformation at the oil−water interface. The morphology of the interfacial nanodroplets was dependent on their location inside the network interfacial layer. The adsorption and deformation of the droplets at the oil−water interface and the formation of network structures, as observed in the present study, provide new, useful fundamental knowledge with potential applications for microencapsulation and emulsion stabilization.
■
INTRODUCTION The adsorption of solid particles at air−water and oil−water interfaces has attracted considerable research attention over the past decade, largely because of the many potential applications in the stabilization of foams and emulsions. The assembly of adsorbed colloidal particles at the surface is expected to provide long-term stability.1,2 Most of the particles selected to produce particle-stabilized emulsions have been synthetic, e.g., polymer latices, silica, metal oxides, and polymeric microgel particles.3−6 Recently, nonsurface-activated CaCO3 nanoparticles have been used to stabilize oil-in-water emulsions through in situ interaction with the anionic surfactant sodium dodecyl sulfate (SDS).7 Moreover, stimuli-sensitive soft microgel particles have been employed as stabilizers for emulsions. These microgel particles are able to adsorb at the oil−water interface and show considerably different behavior from the commonly used Pickering stabilizers, which are solid particles.2,8 Because they are soft and deformable, they present some advantages as emulsifying agents; for example, the deformability of the soft particles at the interface makes it possible to cover a larger interfacial area and form a densely packed compliant shell, which is important for some applications, e.g., in fabricating capsules and preventing coalescence on severe droplet deformation.6,9,10 However, only a few natural materials have been reported to act as particle stabilizers.11−13 Relatively less published research is directly applicable to foods because many of these particles are not permissible in foods. Murray et al.13 © 2013 American Chemical Society
reported on the adsorption behavior of mixtures of surfaceactive polymers and surface-active food-grade particles, such as oil droplets, cellulose, and starch particulates. Recently, some other particles derived from food materials, e.g., starch nanocrystals, flavonoids, nanoparticles, nanoparticle−protein complexes, and lactoferrin nanoparticles, were applied in the formation of the model emulsions.14−19 Proteins are the most important emulsifying agents for making edible oil-in-water emulsions, because of their amphiphilic character and high surface activity. They impart emulsion stability by a combination of electrostatic and steric mechanisms.20,21 One such protein-rich emulsifying agent is the casein micelle that is present in milk.22,23 Casein micelles are considered to be colloidal particles because they consist of aggregated casein proteins with a stabilizing surface layer. κCasein is predominantly on the surface of the casein micelle, βcasein is mostly present in the interior, and αs1- and αs2-caseins are found throughout the micelle structure. It has been reported that the available κ-casein covers about one-third of the surface of the casein micelles and is heterogeneously distributed on the surface, and that the other caseins may share the surface with κ-casein. Casein micelles adsorb at the oil− water interface and then spread along the interface.24,25 The Received: September 9, 2013 Revised: October 29, 2013 Published: October 31, 2013 14403
dx.doi.org/10.1021/la403493y | Langmuir 2013, 29, 14403−14410
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(Nanosizer ZS, Malvern Instruments Ltd., Malvern, Worcestershire, U.K.)), which was similar to the size of the casein micelles in milk.27 However, the average particle size (d32) of the casein micelles is ∼110 nm when the size is determined by static light scattering (Malvern MasterSizer 2000, Malvern Instruments Ltd., Malvern, Worcestershire, U.K.). This indicated that intact casein micelles remained in the dispersion. n-Hexadecane was purchased from Sigma Chemical Co. (St. Louis, MO). All the chemicals used were of analytical grade and were obtained from either BDH Chemicals (BDH Ltd., Poole, U.K.) or Sigma−Aldrich (St. Louis, MO), unless specified otherwise. Preparation of Emulsions. Casein micelle dispersions were prepared by adding micellar casein powder to 10 mM pH 7 phosphate buffer and then stirring for 60 min at room temperature to ensure complete dispersion. The dispersions were adjusted to pH 7.0. Appropriate quantities of n-hexadecane were then mixed with the protein dispersions to give 20% n-hexadecane in the mixtures, which were then homogenized in a microfluidizer (M-110EH; Microfluidics Corporation, Newton, MA) at a pressure of 300 MPa to form the nanoemulsions. Appropriate quantities of n-hexadecane were mixed with the nanoemulsions to give 20% n-hexadecane and 2−16% nanoemulsion in the final emulsions. The mixtures of n-hexadecane and nanoemulsion were then homogenized in a rotor−stator Ultra Turrax homogenizer (10 000 rev/min for 2 min) (Diax 600; Heidolph, Germany) to form the final emulsions. Characterization. A Malvern MasterSizer 2000 (Malvern Instruments Ltd., Malvern, Worcestershire, UK) was used to determine the average diameter (d32) of the emulsion droplets. The relative refractive index (N), i.e., the ratio of the refractive index of the emulsion droplets (1.456) to that of the dispersion medium (1.33), was 1.095. The absorbance value of the emulsion droplets was 0.001. Droplet size measurements are reported as average diameters, d32. The d32 is defined as ∑nidi3/∑nidi2, where ni is the number of particles with diameter di. Mean particle diameters were calculated as the average of duplicate measurements. The protein concentrations of the aqueous phase of the emulsions were characterized as that the emulsions were centrifuged at 45000g for 30 min at 20 °C, the subnatants were then filtered through 0.22 μm filters (Millipore). The protein concentrations of emulsions and the filtrate were determined by the Kjedahl method (1026 Distilling Unit and 1007 Digestor Blorck, Tecator AB, Hoganas, Sweden). The microstructure of the samples was examined using a confocal laser scanning microscopy (CLSM) system (Leica DM6000B, Heidelberg, Germany) with a 63 mm oil immersion objective lens. Nile Blue was used to stain for oil (argon laser with an excitation line of 488 nm) whereas Fast Green was used to stain for protein in the bolus (He−Ne laser with excitation at 633 nm). Samples (1 mL) were stained with 1.0% (wt/vol) Nile Blue and 1.0% (wt/vol) Fast Green (fluorescent dye), placed on a concave confocal microscope slide (Sail; Sailing Medical-Lab Industries Co. Ltd., Suzhou, China), covered with a coverslip, and finally examined with a 63× magnification lens using the microscope. The microstructure of the samples was also studied using transmission electron microscopy (TEM). A sample was mixed with warm melted (35−40 °C) 3% low-temperature-gelling agarose in a 1:1 ratio. This solution was poured on to a microscope slide, allowed to set, and chopped into 1 mm3 cubes. The cubes were put into a bijoux bottle containing 3% glutaraldehyde in 0.2 M sodium cacodylate buffer. This was kept at 5 °C for 24 h. The samples with glutaraldehyde were then rinsed twice with 0.2 M sodium cacodylate buffer over 2 h. The agarose-embedded samples were placed in 1% osmium tetroxide (1 mL) overnight at room temperature. The samples were rinsed twice with distilled water, placed in 1% uranyl acetate (1 mL) for 30 min, and then rinsed twice with distilled water. The dehydration process was carried out at 5 °C in 25% acetone (15 min) and then in 50, 70, and 90% acetone (for 30 min each), followed by 100% acetone (three changes over 90 min). The acetone was replaced with Procure 812 embedding resin, and the samples were put on rollers for 24 h. Sample cubes were placed into an embedding capsule and were cured at 60 °C for 48 h. The embedded samples
redistribution and the spreading of casein micelles at the interface are dependent on the conditions of adsorption, e.g., homogenization pressure, protein concentration, and other emulsifying agents in the system. Murray et al.13 used emulsion droplets coated with protein to strengthen the adsorbed film and to provide enhanced stability for air bubbles, although the emulsion droplets may not have been completely solid. It was considered that the droplet particles had the advantage that they were relatively welldefined in terms of their size, shape, and surface properties. However, the emulsion droplets did not act as a foaming agent because the air bubbles were stabilized by the protein film at the air−water interface. The emulsion droplets associated with the adsorbed protein film under certain conditions, such as at low pH. This association led to the formation of a strong adsorbed film to promote foam stability. In some cases, proteincoated emulsion droplets can become solid particles, e.g., when solid fat such as milk fat or palm oil is used, depending on the environmental temperature. Recently, the use of proteinstabilized emulsion droplets as emulsifying agents to stabilize oil-in-water emulsions was demonstrated in a patent application.26 In this work, we developed a novel stable emulsion system that was stabilized by nanoscale emulsion droplets that were coated with colloidal protein particles (casein micelles). The adsorption and surface properties of both the casein micelles and the casein-micelle-coated emulsion droplets were examined. The arrangement of the casein micelles and caseinmicelle-coated droplets at n-hexadecane droplet interfaces was probed via microscopy. The influence of the particle concentration and the oil/water ratio is discussed.
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EXPERIMENTAL SECTION
Materials. Micellar casein powder was obtained from the American Casein Company (Burlington, NJ). Its composition was as follows: protein 85%, fat 2.1%, ash 9.5%, lactose 3.2% on a dry solids basis, and moisture 4.8%. It was characterized using sodium dodecyl sulfatepolyacrylamide gel electrophoresis (SDS-PAGE) (Figure 1), which showed that only caseins were present. The average particle size of the micellar casein dispersed in 10 mM pH 7 phosphate buffer was found to be ∼200 nm (determined using dynamic light scattering,
Figure 1. SDS-PAGE pattern of micellar casein dispersion. 14404
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were then sectioned to a thickness of 90 nm using a Reichert Ultracut microtome. These sections were mounted on 3 mm copper grids and stained with lead citrate before examination in a Philips transmission electron microscope (NL-5600 MD; Philips, Eindhoven, The Netherlands) at an acceleration voltage of 60 kV. The ζ-potential values of the emulsion samples were measured using a laser Doppler velocimetry and phase analysis light scattering (M3-PALS) technique and a Malvern Zetasizer Nano ZS instrument (Malvern Instruments Ltd.). The samples were diluted to 100 times with Milli-Q water, placed in the electrophoretic mobility cell, and analyzed at a scattering angle of 173° using the Malvern Zetasizer Nano ZS. The effective electric field, E, applied in the measurement cell was between 50 and 150 V depending on the ionic strength of the samples. The overall electrophoretic mobility, μ, was calculated at 20 °C according to eq 1 (for a spherical particle). Y = μω /E
(1) −1
where Y is the particle velocity (m/s) and ω is the frequency (s ) of the applied electric field. The ζ-potential (mV) was then calculated using the Smoluchowski equation: μ = ζε/η
(2)
where ε is the electric permittivity of the solvent (F/m) and η is the solvent viscosity (Pa·s). An individual ζ-potential measurement was calculated from the mean of two runs and the standard deviation of at least 10 readings from an individual sample. A contact angle and surface tension meter (KSV Instruments Ltd., Helsinki, Finland) was used to examine the time evolution of the interfacial tension of protein and protein-coated emulsion droplets via the pendant drop method. The tensiometer operated in a volumecontrolled regime, via continuous measurement of the drop area and volume, approximated by a Laplacian profile. The protein solution or protein-stabilized emulsion was the drop phase against pure nhexadecane. The volume of the drop was about 8 μL. The shape of a drop hanging from a syringe tip was determined from the balance of forces, which included the surface tension of the liquid. The interfacial tension at the liquid interface can be related to the drop shape through eq 3: Δρ × g × R 0 γ= β
Figure 2. (a) Average droplet size (d32) (■) and protein concentration in the aqueous phase (□) of the nanoemulsions formed using micellar casein; (b) droplet size distribution of the nanoemulsion formed at a protein concentration of 40 mg/mL.
(3)
where γ is the interfacial tension, Δρ is the difference in density between fluids at the interface, g is the gravitational constant, R0 is the radius of drop curvature at the apex, and β is the shape factor. The shape factor can be defined through the Young−Laplace equation.
because of flocculation, with the casein micelles bridging between the droplets.28 At higher concentrations, the size of the emulsion droplets remained constant because the excess casein micelles stayed in the aqueous phase. Confocal microscopy of the emulsion formed at a casein micelle concentration of 40 mg/mL indicated a mostly uniform size distribution of small droplets; some droplets appeared to be flocculated (Figure 3). TEM showed clearly that almost all the casein micelles adsorbed on the surface of the emulsion droplets, which had fairly even size distributions (Figure 4). Most of the micelles present at the droplet surface were not intact and appeared to be uneven and spread around the droplet surface (Figure 4c). It is likely that, after adsorption, the casein micelle dissociated and spread along the interface, adjusting its structure to cover the maximum surface area. This resulted in the casein micelles forming a dense and continuous interfacial layer around the emulsion droplets at a relatively low protein concentration. Dissociation and spreading of casein micelles on the surface of fat globules have been observed in previous studies on recombined milk and homogenized milk.25,29 The spreading may have been driven by the thermodynamically favorable changes in the interfacial free energy, which appears to be sufficient to disrupt micelles.30 Another possibility is that homogenization caused disruption of
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RESULTS AND DISCUSSION Formation of Casein-Micelle-Stabilized Nanoemulsion Droplets. Figure 2a shows the diameters of n-hexadecane emulsion droplets stabilized with casein micelles as a function of the total and aqueous phase protein concentrations. The droplet size decreased with increasing protein (mainly casein micelle) concentration in the emulsion system. The average droplet size was about 150 nm when the protein concentration was 40 mg/mL and remained constant above this concentration, indicating the formation of a stable nanoemulsion at a protein concentration of 40 mg/mL; this emulsion showed a narrow droplet size distribution with sizes in the range 50−800 nm (Figure 2b). Very small amounts of protein remained in the aqueous phase (6%, Figure 6), some of the nonadsorbed droplets may interact with adsorbed droplets, resulting in flocs, which may form a dense multilayer with a three-dimensional network at the interface. This multilayer can provide additional stabilization and may play an important role in the stability of particle-stabilized emulsions.1 In a Pickering emulsion, solid particles adsorb and form a thick surface layer, in which the structure of the surface layer is dependent mainly on the particle size and the hydrophobicity, in that the particles need to be partly wetted by both phases.36 Significant deformation of the nanodroplets along the interface of the large droplets, observed in the present study, suggests a different adsorption behavior of soft particles from that of the solid, hard particles used in Pickering emulsion systems, in which the deformation of the solid particles at the interface is negligible. However, the deformation or partial deformation of the particles at the interface has been observed recently for soft particles, such as cross-linked polymeric hydromicrogels formed by NiPAm and BIS, the ligand shell of gold nanoparticles, and copolymer-capped magnetite nanoparticles.6,9,10 Geisel et al.9 intuitively described the deformation of microgel particles at the interface. The equilibrium shape of the microgel particles is given by the balance among the solvations of the hydrogel in the oil and water phases, their interfacial activity, and the internal elasticity of the particles. The particle is stretched radially until the energy gain from the interfacial activity is balanced by the internal elastic deformation of the hydrogel. Compared with these deformable microgel particles and other soft particles, the n-hexadecane nanodroplets used in our study would be softer and more elastic, which could result in more pronounced deformation at the interface, as displayed in Figure 9. The deformation of there nanodroplets is probably driven by a tendency to maximize the interaction of the casein aggregates (adsorbed at nanodroplet surface) with the oil/water interface via the bridging of protein between pairs of interfaces (nanodroplet/droplet). It can be estimated that the extent of deformation is dependent on the relative ratio of the droplet sizes, i.e., nanodroplet:large droplet, the interfacial tension, and the hardness of the interfacial nanodroplets. Furthermore, the deformation may be influenced by the microstructure of the surface layer or the presence of other droplets within the surface layer if a multilayer or three-dimensional network is formed.
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AUTHOR INFORMATION
Corresponding Authors
*A. Ye. E-mail: [email protected]. Tel: 64-6-3505072. *H. Singh. E-mail: [email protected]. Tel: 64-6-3504401. Notes
The authors declare no competing financial interest.
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REFERENCES
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