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Protein fibrils induce emulsion stabilization Jinfeng Peng, Joana Ralfas Simon, Paul Venema, and Erik van der Linden Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.5b04341 • Publication Date (Web): 16 Feb 2016 Downloaded from http://pubs.acs.org on February 17, 2016
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Langmuir
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Protein fibrils induce emulsion stabilization
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Jinfeng Peng, Joana Ralfas Simon, Paul Venema*, Erik van der Linden
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Laboratory of Physics and Physical Chemistry of Foods, Department of Agrotechnology and
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Food Sciences, Wageningen University, P.O. BOX 17, 6700 AA, Wageningen, The Netherlands
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Keywords: protein fibrils, emulsion, stable clusters, depletion stabilization
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ABSTRACT
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The behavior of an oil-in-water emulsion was studied in the presence of protein fibrils for a
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wide range of fibril concentrations by using rheology, diffusing wave spectroscopy and confocal
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laser scanning microscopy. Results showed that above a minimum fibril concentration, depletion
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flocculation occurred, leading to oil droplet aggregation and enhanced creaming of the emulsion.
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Upon further increasing the concentration of the protein fibrils, the emulsions were stabilized. In
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this stable regime both aggregates of droplets and single droplets are present and these
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aggregates are smaller than the aggregates in the flocculated emulsion samples at the lower fibril
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concentrations. The size of the droplet aggregates in the stabilized emulsions is independent of
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fibril concentration. In addition, the droplet aggregation was reversible upon dilution both by a
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pH 2 HCl solution and by a fibril solution at the same concentration. The viscosity of the
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emulsions containing fibrils was comparable to that of the pure fibril solution. Neither fibril
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networks nor droplet gel networks were observed in our study. The stabilization mechanism of
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emulsions containing long protein fibrils at high protein fibril concentrations points towards the
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mechanism of a kinetic stabilization.
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INTRODUCTION
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Proteins have the ability to form various structures under different conditions. The
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morphologies of these structures range from linear to spherical. One of the linear structures is
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protein fibrils that can be formed using a wide range of proteins, such as, egg white proteins
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(ovalbumin1 and lysozyme2), soy protein3, whey protein4 and pea protein5, when the appropriate
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conditions are used. For example, when heating β-Lactoglobulin (β-lg) at low pH and low ionic
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strength, the protein is first hydrolyzed into small peptides, and part of these peptides assemble
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into fibrils.6, 7, 8 These fibrils typically have a diameter in the order of nanometers, a length in the
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order of micrometers,9 and a persistence length around a micrometer.10, 11 Their high aspect ratio
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and rod-like structure leads to a high excluded volume, 12 making them interesting candidates as
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additives to colloidal dispersions.
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The addition of non-adsorbing polymers to colloidal dispersions, can changes the stability of
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the dispersion and has been frequently studied due to its fundamental aspects and wide range of
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applications in different fields.13 Above a minimum polymer concentration, polymers can induce
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depletion flocculation of the colloidal particles leading to phase separation.14, 15 The flocculation
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rate increased with increasing polymer concentration, until reaching a critical concentration,
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above which the rate of flocculation decreased with increasing polymer concentration. At high
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enough polymer concentrations, the dispersion becomes stable. This phenomenon is known as
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depletion stabilization.16 Rods have been found to induce a rich phase behavior in colloidal
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dispersions.15,
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emulsions.21 They suggested that the flocculation at low fibril concentrations was induced by
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Blijdensteijn et al. studied the behavior of rod-like protein fibrils in
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depletion interaction. Here we investigate the stabilization mechanism of emulsions at high fibril
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concentrations, where the rod length and oil droplet diameter are comparable. Earlier studies on
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depletion interaction mainly focused on relatively low concentration of rods, and the size of rods
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being much smaller than that of the particles.15, 22, 23, 24, 25, 26, 27, 28 We use rheology, diffusing
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wave spectroscopy and confocal laser scanning microscopy as experimental techniques.
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EXPERIMENTAL SECTION
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Whey protein isolate (WPI) solution preparation
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WPI powder (Bipro, Davisco, Lot # JE 198-1-420, USA ) was dissolved in HCl solution of
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pH 2 to prepare a stock solution of 10 wt % WPI. The solution was left overnight in a cold room
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(4 °C) while mildly stirring using a magnetic stirring bar. After dissolution, the pH of the protein
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solution was adjusted to pH 2 with a 6 M HCl solution. Subsequently, the protein solution was
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filtered through a filter (FD 30/0.45 µm Ca-S from Schleicher & Schuell). The protein
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concentration of the stock solution was determined using a UV spectrophotometer (Cary 50 Bio,
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Varian) at a wavelength of 278 nm, and a calibration curve determined from WPI-solutions at
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known concentrations. The WPI stock solution was stored at 4°C for further use in fibril
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formation and emulsion preparation.
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Fibril formation
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The WPI stock solution was diluted to a protein concentration of 4 wt % with a HCl solution
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at pH 2. The samples were placed in a heating block (RT 15 Power, IKA-WERKE) and stirred at
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80°C for 22 hours to form fibrils. After this step the fibril solution was cooled down in ice-water
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and afterwards stored at 4°C. The fibril concentration indicated throughout this paper refers to
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the protein concentration before heating.
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We note that for the preparation of the emulsions with 3.25 wt % fibrils we had to use a stock
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solution of fibrils prepared at 4.5 wt % of protein. This does not affect the average fibril length.4
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Oil-in-water (o/w) emulsion preparation
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WPI was used as an emulsifier to prepare a 40% (v/v) o/w emulsion. The emulsion was
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prepared in two steps. First, sunflower oil (Reddy, The Netherlands, obtained from a local
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supermarket) was mixed with a 1 wt % WPI solution using a high speed blender ( Ultra Turrax,
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T25 digital , IKA), at a speed of 9000 RPM for 5 minutes. Subsequently, the emulsion was
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homogenized by 5 passes through a homogenizer (Labscope, Delta instruments, The
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Netherlands) at a pressure of 70 bar. The emulsion was stored at 4 °C before use and all
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experiments were performed within 12 hours after storage.
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Sample preparation
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A range of concentrations of protein fibrils were prepared by diluting the 4 wt % protein fibril
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solution and the emulsion was added to those fibril solutions. The final volume percentage of
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emulsion droplets in the mixture was 10 % (v/v). The concentration of protein fibrils in the
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mixture were : 0 (no fibrils added), 0.001, 0.005, 0.01, 0.05, 0.1, 0.5, 1, 1.5, 2, 2.25, 2.5 , 2.75, 3
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or 3.25 wt %. The emulsion and protein fibril solution were mixed by mild agitation for 10 s.
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Characterization of protein fibrils and emulsion
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Length distribution of protein fibrils
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The length distribution of the fibrils was determined using flow-induced birefringence. The
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decay curves of the flow-induced birefringence after the cessation of flow were measured with a
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strain-controlled ARES rheometer (Rheometrics Scientific) equipped with a modified optical
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analysis module.29 From these decay curves the length distribution of the fibrils were
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determined.30
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Transmission electron microscopy (TEM)
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TEM was used to image the protein fibrils. After a 100-fold dilution a droplet of the sample
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was placed on a carbon film supported by a copper grid. After 15 s, the droplet was brought into
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contact with a filter paper to remove most of the water. Subsequently, a droplet of 2% uranyl
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acetate was added and removed after 15 s by another filter paper. Electron micrographs were
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taken using a transmission electron microscope (JOEL JEM-1011, Tokyo, Japan).
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Size distribution of oil droplets
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The size distribution of oil droplets was measured using multi-angle static laser light scattering
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(Mastersizer MS2000, Malvern Instruments). The average diameter of the oil droplets, D[3,2], was
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around 1.9 ± 0.2 µm for all batches.
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Characterization of emulsion stability
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Visual observation
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Samples were placed at room temperature (20 °C) and images were taken directly after
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preparation (0 days) and after 1 day and 7 days.
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Creaming stability
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The emulsions were placed in glass tubes and monitored over time at 20 ᵒC. The
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backscattering intensity of laser light as a function of height was measured (Turbiscan, MA
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2000). The intensity profile was measured at times 0, 1 and 7 days. Interpretation of the data is
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based on an analysis described elsewhere.21, 31
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Confocal laser scanning microscopy (CLSM)
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The microstructure of the cream layer of samples after 1 day and 7 days was imaged using
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CLSM (Zeiss 200 m Axiovert, Thornwood, NY, USA). Samples were placed in a chamber
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prepared from a microscope glass slide (Menzel-glaser, 50 mm, Thermo Scientific,
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Braunschweig, Germany) and a cover slip (Menzel-Glaser, 15 mm × 15 mm, Thermo Scientific,
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Braunschweig, Germany). Images were made using an oil immersion objective (Plan-
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Apochromat, 100x/1.46 oil, Zeiss, Germany). One drop of 0.2 wt % Nile blue (Merck, Germany)
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solution was added to the samples before imaging. Sample imaging was performed at an
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excitation wavelength of 488 nm and an emission wavelength of 525 nm.
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Viscosity
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The viscosity of the samples was measured on a stress controlled rheometer (MCR302, Anton
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Paar), using a concentric cylinder geometry (CC17/T1/S-SN38492). A solvent trap was used to
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prevent evaporation. The samples were pre-sheared at a shear rate of 30 s-1. Subsequently, the
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viscosity was monitored by increasing the shear rate from 0.01 to 500 s-1. Measurements were
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performed at 20 °C.
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Strain sweep
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A strain sweep was performed on a stress controlled rheometer (MCR501, Anton Paar), using
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a concentric cylinder geometry (CC17/T1-SN3951). A solvent trap was used to prevent
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evaporation. The strain sweep was carried out at 20 °C, at frequency of 0.1 Hz varying strain
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from 0.01% to 100%.
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Diffusing wave spectroscopy (DWS)
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In our study, DWS is used to monitor the diffusive motion of emulsion droplets at various
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fibril concentrations. Dispersions of only fibrils are completely transparent, therefore, DWS is a
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suitable technique to monitor the emulsion droplets.
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The samples were analyzed directly after mixing using DWS in transmission mode. In DWS,
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the intensity autocorrelation function g2(t) of the multiple scattered light is measured. Following
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ref 32 g2(t) is related to the mobility of the emulsion droplets as
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∞ s g 2 (t ) − 1 = β ∫ P( s ) exp − 1 3 k 02 < ∆r 2 (t ) > * ds l 0
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where it follows from the Siegert relation that
g 2 (t ) − 1 = β g1 (t )
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(2)
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Here β is a constant determined by the collection optics, P(s) is the path length distribution
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function of paths of length s, k0 = 2πn/λ, with n the refractive index of the solvent and λ the
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wavelength of the laser, is the mean square displacement of the droplets , and l* is the
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photo transport mean free path.
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The motion of the emulsion droplets, expressed by their mean square displacement (MSD)
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can be obtained by numerically inverting the correlation functions and using Equation
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Lk 1. Since it is convenient to work in scaled variables we define Q = ∗ 0 < ∆r 2 (t ) > . L refers to l
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the optical path length through the sample cell. In a purely viscous solution, the oil droplets
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would undergo Brownian motion (unhindered diffusive colloidal motion) and Q2 would vary
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linearly with time Q2 ~ Dt, where D is the diffusion constant. In a viscoelastic medium the short
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time behavior of Q2 can generally be described by a power law < Q 2 (t ) > ≈ Dt p where D is the
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diffusion constant, p is a measure for the viscoelastic properties of the matrix. When p ≤
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matrix behaves like a medium where G’≥ G’’, when ½ < p
