Food &Function

PAPER

Cite this: DOI: 10.1039/c6fo01027j

Received 9th July 2016,Accepted 24th September 2016

DOI: 10.1039/c6fo01027j

www.rsc.org/foodfunction

High internal phase emulsions stabilized solely bywhey protein isolate-low methoxyl pectincomplexes: effect of pH and polymerconcentration†

Wahyu Wijaya,a,b Paul Van der Meeren,a Christofora Hanny Wijayac andAshok R. Patel*b

In recent years, there has been significant progress in edible emulsion technology especially with

respect to creating and stabilizing surfactant-free emulsion systems for food applications. In this paper,

we demonstrate the fabrication of high internal phase emulsions (HIPE) (φoil = 0.82) stabilized using

colloidal complexes of non-gelling biopolymers (at concentrations as low as 0.3 wt%). The colloidal

complexes were pre-formed by combining whey protein isolate (WPI) and low-methoxyl pectin (LMP) at

three different pH values (i.e. pH 3.5, 4.5, 5.5) and used further for fabricating stable HIPEs. In addition to

the effect of pH, the influence of total biopolymer concentration on the formation and properties of

HIPEs was also evaluated. Depending on the total concentration of biopolymers used, the WPI–LMP

complexes (formed at pH 4.5) showed a Z-average diameter in the range of 250–350 nm. It was found

that the formation of HIPEs was strongly influenced by the pH of the colloidal complexes. At a pH close

to the isoelectric point of WPI (≈pH 4.8) and WPI–LMP complexes (≈pH 3.4), severe aggregation of col-

loidal particles occurred, resulting in poor formation and stability of HIPEs. On comparing the stabilization

behaviour of the complexes with the uncomplexed protein, it was noticed that the former provided com-

paratively better stabilization to the HIPEs against coalescence at pH 4.5 and 5.5. Based on the rheological

data (low amplitude oscillatory shear rheology and flow measurements), all HIPE samples showed visco-

elastic and shear-thinning behaviour. We believe that such viscoelastic gel-like systems could find poten-

tial commercial applications in the development of label-friendly novel food products with interesting

textures.

Introduction

A colloidal system is defined as a dispersion where discreteparticles (droplets, bubbles, solid particles, etc.) of one phaseare homogeneously dispersed in a continuous phase ormedium, and the dispersed particles are typically in the sizerange of nanometers to tens of microns.1 Complex colloidal

systems are a sub-category of colloidal systems which haverecently gained a lot of attention from researchers working inmultidisciplinary fields. Although there is no clear definitionof this broad sub-category, they include colloidal systems thatcontain more than one type of colloids (e.g. emulsion gels,2

foamulsions,3 suspoemulsions,4 etc.) or consist of more thantwo phases (e.g. double emulsions where oil and water are dis-tributed into multiple phases)5 or they may have an unusualphase distribution (e.g. High Internal Phase Emulsions,HIPEs, where the volume fraction of the dispersed phase isabove the close packing volume)6 or the viscoelasticity of oneor more phases is altered through solidification or immobili-zation (structured emulsions).7

In the past few years, complex colloids such as HIPEs havebeen an attractive subject of investigation due to their widerange of applications particularly in the structuring of edibleformulations.6,8 HIPEs are emulsified systems with an internalphase volume fraction of at least 0.74 (φdisp ≥ 0.74), which is themaximum packing density of monodispersed hard spheres.9

†Electronic supplementary information (ESI) available. See DOI: 10.1039/c6fo01027j

aParticle and Interfacial Technology Group, Department of Applied Analytical and

Physical Chemistry, Faculty of Bioscience Engineering, Ghent University, Coupure

Links 653, B-9000 Gent, Belgium. E-mail: wahyu.wijaya@ugent.bebVandemoortele Centre for Lipid Science and Technology, Laboratory of Food

Technology and Engineering, Department of Food Safety and Quality, Faculty of

Bioscience Engineering, Ghent University, Coupure links 653, 9000 Gent, Belgium.

E-mail: ashok2510@gmail.comcDepartment of Food Science and Technology, Faculty of Agricultural Technology,

Bogor Agricultural University, Kampus IPB Darmaga Bogor 1668 PO. Box. 220,

Indonesia

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It is well-known that various food products use concen-trated emulsions or HIPEs in their formulations (e.g. mayon-naise and salad dressings).10 These systems are generallystabilized using low-molecular weight surfactants which areused at relatively high concentrations to prevent serious stabi-lity issues such as coalescence and phase inversion.11 One ofthe most pressing needs in the food manufacturing industrycurrently is to provide well-informed consumers with theoption of food products that are label friendly (created usingnatural ingredients). Accordingly, most food manufacturersare currently interested in investigating the possibility ofreplacing synthetic surfactants and small molecular weightemulsifiers with biopolymers such as proteins and poly-saccharides as natural sources.12

Although proteins are excellent emulsifiers (due to theirinherent amphiphilicity), their stabilizing functionality can begreatly improved by combining them with polysaccharidesto create soluble conjugates or precipitated colloidalcomplexes.13,14

With respect to colloidal complexes, many studies focusingon the emulsion stabilization properties of protein–poly-saccharide electrostatic complexes have already been reportedsuch as whey protein isolate–carrageenan,11 whey protein–pectin,15 sodium caseinate–chitosan,16 and ovalbumin–pectin,17 confirming that this is a highly active area ofresearch. However, unfortunately majority of these studieshave been performed on rather dilute emulsion systems (φoil ≈0.1 or 0.2) while their functionality in stabilizing concentratedemulsions has rarely been reported and to our best knowledgedirect fabrication of high internal phase oil-in-water emulsionswith protein–polysaccharide electrostatic complexes has notbeen explored to date.

Thus, the main aim of the current study was to investigatethe possibility of using protein–polysaccharide complexes tofabricate and stabilize HIPEs. To achieve our goal, wheyprotein isolate (WPI) and low methoxyl pectin (LMP) were usedas representative models of non-gelling proteins and anionicpolysaccharides to first produce colloidal complexes(via electrostatic or charge-induced interactions) followed byinvestigating their functionality in stabilizing water continuousHIPEs (φoil = 0.82).

WPI is commonly used as a natural emulsifier in foods tofacilitate the formation and stabilization of oil-in-water emul-sions.18 It is a mixture of different globular proteins, withβ-lactoglobulin being the most dominant fraction followed byα-lactalbumin.19 Unlike gelatin which has gelling propertiessimilar to other hydrocolloids,20 native WPI is typically notknown to form an aqueous gel without heat21 or enzymatictreatment (e.g. transglutaminase),22 or addition of divalentsalts (e.g. calcium chloride).23 Pectin is a substance found inthe middle lamella and primary cell walls of different planttissues. It consists of a linear backbone of (1,4) α-D-galac-turonic acid residues that are partially esterified with methanoland depending on the degree of methylation (molar ratio ofmethanol to galacturonic acid) are categorized as low or highmethoxyl pectins.24

In this work, WPI–LMP colloidal complexes were pre-formed by pH adjustment followed by fabrication of HIPEsthrough high-shear homogenization of the oil phase with theaqueous dispersion of colloidal particles. Hereby, the effect ofpH, biopolymer concentration and proportion of individualbiopolymers on the formation and properties of HIPEs werestudied. The microstructure and bulk properties of emulsionswere further investigated using advanced microscopy (cryo-SEM) and small deformation studies (oscillatory rheology andflow measurements), respectively.

Experimental sectionMaterials

A minimally heat-treated whey protein isolate (WPI), enrichedin β-lactoglobulin (approx. 85% of total protein), was obtainedfrom Davisco Foods International, Inc. (Le Sueur, MN, USA).Unipectine OB700 – a low methoxy apple pectin with a DEbetween 33 and 38% was received from Cargill R&D Vivoorde,Belgium. Sodium azide, sodium chloride, and HCl wereobtained from Sigma Aldrich. NaOH was bought from VWRChemicals and refined sunflower oil was obtained from a localsupermarket. Milli-Q water was used throughout the study. Allmaterials were used without further purification or modifi-cation, and all samples were formulated and reported on aweight-by-weight basis (g g−1).

Complex formation and emulsion preparation

Preparation of stock solutions. WPI and LMP powders wereweighed into separate beakers at 5% and 2.5% respectively in100 mL Milli-Q water containing 0.02% sodium azide. Thesedispersions were then stirred continuously at room tempera-ture (22–25 °C) until complete dissolution was obtained. Thesolutions were then stored overnight at 5 °C to ensure com-plete hydration of biopolymers.

Complex formation. WPI and LMP stock solutions werediluted and mixed together to obtain a concentration of1.0 : 0.5% in the aqueous phase. Afterwards, the solutions wereadjusted to pH 3.5, 4.5, and 5.5 with dilute hydrochloric acid.

Emulsion preparation. HIPEs were prepared by homo-genizing sunflower oil (concentration of 80%, φoil = 0.82) andan appropriately diluted aqueous phase (20%) containingcolloidal complexes formed at pH 3.5, 4.5 and 5.5 to obtainWPI : LMP concentrations of 1.0 : 0.5%, 1.5 : 0.75%, 2.0 : 1.0%and 2.5 : 1.25% in the aqueous phase. All emulsions werehomogenized for 2 min at 13 500 rpm using an UltraTurraxIKA® T25 Basic (IKA-Werke GmbH & Co. KG, Germany)equipped with a dispersing rotor S25KV-25G. The resultingself-standing emulsion gels were then stored at 5 °C prior toanalysis on the following day. For comparison, emulsions withonly WPI were also processed in a similar manner.

Particle characterization

Charge titration. WPI (1.0%), LMP (0.5%) and the WPI : LMPdispersion (1 : 0.5%) in the 200 mL final solution were titrated

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with 1 N HCl from pH 7.0 down to pH 1.5. The streamingpotential signal was recorded by using a Charge Analyser II(Rank Brothers Ltd, England) equipped with a streamingpotential cell.

Turbidity measurements. The turbidity of the dispersionswas measured by using an UV-Vis spectrophotometer(UV 1205, Shimadzu FS 689), at λ = 600 nm.

Z-Average diameter. Measurements for particle size distri-bution were carried out using photon correlation spectroscopy(PCS) with a Spectrometer 100 SM (Malvern Instruments,Malvern, UK) equipped with a 15 mW He–He laserK7032 multi 8-bit correlator (Malvern Instruments, Malvern,UK). Analyses were carried out at a scattering angle of 150° at25 °C with 50 or 100 μm aperture. The Z-average mean dia-meter was obtained by cumulant analysis. All measurementswere done in triplicate.

Surface tension. The surface tension of WPI–LMP disper-sions was measured by the Wilhelmy Plate method. Themeasurement was performed at room temperature. Prior tothe experiments, all glassware in contact with the biopolymersolutions was properly cleaned with a concentrated chromicacid solution and abundantly rinsed with Milli-Q water.

Contact angle measurements. A homogeneous WPI : LMPcolloidal dispersion was poured into a pre-cleaned glasschamber. The dispersion was then dried at controlled roomtemperature (22 ± 2 °C). These dried WPI–LMP complexes wereground and pressed to produce a film. The contact angle (θw)of a sessile drop of water (10 µL) on the WPI–LMP film wasmeasured using a drop shape analysis system DSA 10 Mk2(Kruss GmbH, Germany). Contact angles were determinedfrom approximating the contour of the imaged droplets with acircle fitting.

Specific surface area calculation. The total surface area ofprotein per unit volume of emulsion was calculated by thefollowing equation:

SSA ¼ 6D½3;2� � ρoil

where SSA is the specific surface area (m2 kg−1), D[3,2] is theSauter mean diameter of the emulsion droplets (m), and ρoil isthe sunflower oil density (920 kg m−3). The SSA is influencedby free versus adsorbed protein (not all protein is adsorbed).

Emulsion characterization

Droplet size distribution. The volume-weighted averagedroplet size (D[4,3]) of emulsions was determined using aMasterSizer 3000 (Malvern Instruments Ltd, Malvern,Worcestershire, UK) equipped with a wet sample dispersionunit (Malvern Hydro MV, UK). Before measurement allsamples were diluted 20 times with the same pH of the solu-tion. In the sample port, the samples were dispersed in mQwater at 1500 rpm until an obscuration of 2–6% was obtained.The background and sample integration times were 20 and10 seconds, respectively. The optical properties were definedas refractive index 1.47 (sunflower oil) and 1.330 (dispersantwater) and absorption index 0.01, with the normal instrument

calculation sensitivity and general purpose spherical particleshape selected. Results were calculated with the MastersizerVersion 5.54 software (Malvern, UK) to obtain the particle sizedistributions and the volume-weighted average droplet size(D4,3). All measurements were done in triplicate.

Microstructure studies. Optical, and cryo-scanning electronmicroscopy techniques were utilized to study the microstruc-ture of the samples. Optical microscopy was carried out on aLeica DM2500 microscope (Leica Microsystems, Belgium). Forcryo-SEM, samples of the emulsions were placed in the slots ofa stub, plunge-frozen in liquid nitrogen, and transferred intothe cryo-preparation chamber (PP3010 T cryo-SEM preparationsystem, Quorum Technologies, UK), where they were freeze-fractured, sublimated (only for emulsions) and subsequentlysputter-coated with Pt and examined with a JEOL JSM 7100FSEM (JEOL Ltd, Tokyo, Japan).

Rheological measurements

Viscosity. The viscosity of the WPI–LMP dispersions wasmeasured on an advanced rheometer AR 2000ex (TAInstruments, USA). The concentric cylinder geometry wasequipped with a standard cup radius of 15 mm, and config-ured with a DIN Rotor (radius of 14 mm and height of42 mm). Viscosity was measured at a range of shear rates of0–20 s−1 at 25 °C. A shear rate of 10 ( s−1) was then used todisplay the viscosity of the WPI–LMP dispersions.

Small amplitude oscillatory shear rheology. Dynamic rheo-logical measurements were carried out on an advanced rheo-meter AR 2000ex (TA Instruments, USA) equipped with aPeltier system for temperature control. A parallel plate cross-hatched geometry of 40 mm diameter was used and the geo-metry gap was set at 1000 μm. A range of experiments includ-ing an amplitude sweep (stress = 0.1–1000 Pa, frequency = 1Hz), and frequency sweeps (0.1–10 Hz, oscillatory stress = 2 Pa)were carried out at 5 °C. For flow measurements, samples weresubjected to increasing shear rates (1 and 20 s−1) at 5 °C. Inthe amplitude sweep measurements, G′LVR was obtained bycalculating the mean of G′ before the critical yielding point ofthe G′, while oscillatory yield stress (σy) was the crossover pointbetween G″ and G′.

Stability studies

HIPE stability. The stability of the HIPEs was evaluated bymeasuring the O/W phase separation under centrifugal forcewith an analytical photo-centrifuge (LUMiFuge 116, L. U. M.GmbH, Germany). In this study, 2 g of the sample in a 5 mLglass tube (≈20 mm measurement region) was centrifuged at2000 rpm (540g) at 25 °C for 1 h. During the centrifugation,the near-infrared sensor scanned the sample cells over thetotal length. A charge coupled device (CCD) line sensorreceived light transmitted through the sample, which showeda pattern of light flux as a function of the radial position,giving a transmission profile of the sample at a given time,from which emulsion instability, such as creaming, sedimen-tation, and coalescence could be detected. Quantitative datawere obtained by integration of the transmission profiles

Food & Function Paper

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between the 106–115 mm radial positions from the rotationcenter. The total integrated area was plotted against the cen-trifugation time. The slope of this graph was used to expressthe separation rate and thus the stability, with its lower valueindicating a better creaming stability. The slope was directlyobtained by using SEPView 4.0 Lumifuge software, through thecalculation of linear regression during the initial period ofsteep rise of transmission before the plateau was reached,i.e. between 0 s and 1000 s for unstable emulsions andbetween 0 s and 3600 s for stable emulsions.

Ultracentrifugation induced-coalescence. Coalescence wasstudied by ultracentrifugation (Beckman L7–55, USA) with aSW 40 Ti rotor. Four grams of samples were placed in 6 mLultracentrifuge tubes. The ultracentrifuge was operated at25 °C, 20 000 rpm (70 952g) for 60 min. After centrifugation,some of the emulsions were separated into different phases,i.e. an oil layer on the top, a cream layer at the middle and aserum layer at the bottom of the test tube.

Data analysis

All experiments were carried out at least in duplicate usingfreshly prepared samples. All data analyses were done usingMS-Excel 2010 and Origin 8.0. The results are reported asmeans and standard deviations.

Results and discussionFormation and morphology of HIPEs

WPI : LMP colloidal complexes were pre-formed by pH adjust-ment of aqueous mixtures (pH = 3.5, 4.5, and 5.5). Theseaqueous dispersions containing complexes of WPI : LMP werethen used as the aqueous phase and homogenized togetherwith sunflower oil (oil/aqueous phase mass ratio = 4 : 1) toobtain HIPEs. Homogenization was done under high shear(13 500 rpm) of mixing to force adsorption of the complexeson the surface of the oil droplets for efficient coverage of theoil–water interfaces. Due to the high volume of the dispersedphase, the closely packed oil droplets provided a structuralframework that held the emulsion together with the resultantformation of a self-standing elastic gel-like structure. For com-parison, HIPEs stabilized by uncomplexed WPI were also pre-pared using a similar protocol to that described above.

The representative pictures of the macro- and microstruc-ture of HIPEs stabilized by uncomplexed WPI and WPI–LMPcomplexes are shown in Fig. 1 and 2 respectively. As indicatedby the microscopy images, the pH strongly influences bothcomplex formation and subsequent stabilization of HIPEs.HIPEs prepared using uncomplexed WPI (at all pH values)showed poor stability, indicated by prominent coalescence.The instability because of the large oil droplet size was evengreater at pH 4.5 due to severe aggregation of WPI particles atits pI (≈pH 4.8) as observed in Table 1. Likewise, HIPEs pre-pared using WPI–LMP complexes at pH 3.5 also showedinstability due to the aggregation of the complexes at their pI(≈pH 3.4) (Table 1). The determination of the pI of WPI and

complexes based on charge titration and turbidity analysis isshown in Fig. S1 (ESI†). On the other hand, HIPEs preparedusing complexes at pH 4.5 and 5.5 showed the finest appear-ance as confirmed from the microscopy images where the oildroplets were seen to be closely packed together with theabsence of any coalescence at the microscale or oiling-off at

Fig. 1 Photographs and optical microscopy images (100× magnifi-cation) of HIPEs stabilized by WPI (1.0%) containing 80% sunflower oil.Bars are 20 μm. All images were taken one day after preparation.

Fig. 2 Photographs and optical microscopy images (100× magnifi-cation) of HIPEs stabilized by complexes (1.0 : 0.5%) containing 80%sunflower oil. Bars are 20 μm All images were taken one day afterpreparation.

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the macroscale level (over 5 months of storage). In addition,the viscoelasticity of the HIPEs also contributed to their easyspreadability without any signs of emulsion breakage and oilexpulsion under shear encountered during spreading.

We also examined the effect of the total polymer concen-tration as well as the ratios of the monocomponents (WPI andLMP) on the formation of complexes and HIPEs (Fig. 3). Fourdifferent concentrations of WPI : LMP were evaluated at pH4.5. As expected, an increase in the total concentrationresulted in smaller droplet sizes of the dispersed oil phase.The highest concentration investigated in this study was2.5 : 1.25% of WPI : LMP in the aqueous phase. Above this con-centration solubility limitations made it difficult to fully dis-perse and hydrate the polymers.

In general, the pH of WPI and complex formation stronglyinfluenced the macro- and microstructure of the HIPE. Thecoalescence observed in the emulsions stabilized by WPIformed around the pI of WPI (pH 4.5–5.5)25 could be attribu-ted to the formation of larger protein aggregates. However, atpH 3.5, where WPI particles in the nanoscale range wereformed (Table 1), there was also evidence of extensive dropletcoalescence (Fig. 1). This is similar to the findings reported forsodium caseinate-stabilized emulsions at pH 3 where the inter-action between the positively charged protein particles resulted

in poor adsorption onto the oil interfaces.26 In the case ofWPI–LMP complexes, coalescence was only observed at pH 3.5and the consistency of stable emulsions prepared at 4.5 and5.5 could be tuned to smooth (mayonnaise-like) and firm(margarine-like) spreadable textures depending on the concen-tration of complexes used in emulsion preparation.

A speculative mechanism (based on cryo-SEM images)involved in the stabilization of emulsions is shown in Fig. 4. InHIPEs stabilized by WPI, oil droplets are proposed to becovered by a molecular layer of globular protein,β-lactoglobulin being the major protein in WPI consisting of162 amino acid residues and a molecular mass of about18.3 kDa.27,28 These globular proteins form a condensed filmaround the oil droplets resulting in the stabilization of HIPEs.In addition, larger particulates could also be formed duringpH adjustment resulting in the formation of spherical clus-tered particles. These particles can also anchor onto oil–waterinterfaces and contribute to the stabilization of HIPEs. Foremulsions stabilized by complexes, the stabilization isachieved through a combination of an interpolyelectrolytenetwork (comprising of pectin chains linked together with clus-tered protein particulates) between the droplets and an inter-mixed layer of WPI–LMP complexes. Similar types of structureshave been reported by Le and Turgeon (2013) where they suggestthat intramolecular interactions between the polysaccharidechains and the protein aggregates can occur at pH ≥ pI ofprotein for β-lactoglobulin–xanthan gum combinations.29,30

Particle size and droplet size measurement

The Z-average diameter of the complexes contributes to theHIPE formation and the oil droplet size. Table 1 presents theeffect of pH on the Z-average diameter of WPI : LMP complexesand on the volume weighted mean diameter of HIPEs. Forcomparison, the data of WPI are also given. WPI at pH 5.5 wasslightly aggregated (≈1 μm) as the pH was in the range of thepI of WPI, which was also shown by the slightly white appear-ance of the WPI solution indicating an increase of turbidity in

Table 1 Z-Average diameter of WPI (1.0%) and complexes (1.0 : 0.5%),and D[4,3] of HIPEs stabilized by WPI and complexes at differentpH values

pH

Z-Average (nm) D[4,3] (µm)

WPI Complexes WPI Complexes

3.5 153 ± 3 9171 ± 2340 16.10 ± 0.26 19.00 ± 0.784.5 35 413 ± 5000 222 ± 12 44.09 ± 2.01 14.83 ± 0.255.5 1049 ± 101 440 ± 4 16.03 ± 0.57 12.10 ± 0.61

Fig. 3 Optical microscopy images (200× magnification) of fresh HIPEsstabilized by complexes with concentrations of 1.0 : 0.5% (A), 1.5 : 0.75%(B), 2.0 : 1.0% (C), 2.5 : 1.25% (D) at pH 4.5. Bars are 20 μm. All imageswere taken one day after preparation.

Fig. 4 Cryo-SEM images and schematic of HIPE stabilized by WPI par-ticles (top) and complexes (below) formed at pH 5.5. Bars are 1 µm.

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the WPI solution. The effect was even greater at pH 4.5 as thisvalue is quite close to the pI of WPI (≈pH 4.8), in which thescattered intensity of the WPI solution showed a maximumwith a milky appearance and some precipitation. Thus, the oildroplets at this pH had a very large diameter (≈44 µm) asexpected. At pH 3.5, which was far from the pI of WPI, the col-loidal dispersion was quite stable with a particle size in thenanoscale range (≈153 nm), which is in line with the clearsolution appearance. In addition, emulsions formed by WPI atpH 3.5 and 5.5 showed similar droplet sizes (≈16 µm) eventhough the sizes of the WPI particles were different underthese two pH conditions.

In contrast, the Z-average diameter of the complexes at pH3.5 was larger than 5 µm due to the proximity to the pI of thecomplexes as confirmed by charge titration and turbidity data(ESI, Fig. S1†). At pH 4.5 and 5.5, stable and fine complexeswere formed with particle sizes in the nanoscale range. At thesepH values, the emulsions also showed a smaller droplet sizecompared to HIPEs stabilized by WPI, suggesting the stabilizingrole played by the presence of pectin in the complexes.31

Increasing the concentration of WPI : LMP in the complexesled to an increase in the viscosity of the dispersion and anincrease in the size of the particles (Table 2). Interestingly,although the particle size of the complexes increased with theincrease of biopolymer concentration, the emulsions stabilizedwith these dispersions showed a decrease in the size of the oildroplets. This suggests that the size of the particles alone isnot the deciding factor for emulsion stabilization; it is ratherthe total concentration of particles which can provide anefficient coverage of the interfacial boundaries.32

As seen from Fig. 5, the particle size distributions of theHIPEs are represented by bimodal and trimodal curves. Theminor peaks on the left which were observed at all pH valuesrepresent the unadsorbed particles in the bulk phase. HIPEsstabilized by WPI at pH 3.5 showed a distribution with themajor peak position shifted slightly to the left towards asmaller droplet size than that stabilized by complexes. In con-trast, at pH 4.5, larger droplet sizes and a broader distributionrange were observed for WPI stabilized emulsions as comparedto the emulsions stabilized by complexes. Both HIPEs stabi-lized by WPI or complexes at pH 5.5 showed a narrow distri-bution range with the HIPEs stabilized by complexes showingcomparatively smaller droplet sizes. The increase in the totalbiopolymer concentration led to a decrease in the droplet sizeas discussed above.

On comparing the non-aggregated systems (i.e. HIPEsstabilized by WPI at pH 3.5 and 5.5 and HIPEs stabilized bycomplexes at pH 4.5 and 5.5), we found that a higher pHresulted in a smaller oil droplet size for WPI and complexes,even though the particles were larger (Table 1). It can be specu-lated that at low pH, the electrostatic repulsion between thehighly charged protein molecules was strong enough to reducethe hydrophobic attraction among particles at the interfacewhich potentially reduces their alignment and intermolecularinteraction on surfaces, thus leading to a larger droplet sizedue to less protein coverage.33

The protein coverage on the interface was also affected bythe total protein concentration, a higher concentration result-ing in a smaller droplet size. Increasing the total biopolymerconcentration reduced the volume weighted mean diameter ofthe oil droplets resulting in a larger surface area per unitvolume of emulsion.34 In most cases, D[3,2] or the volume/surface average or the Sauter mean diameter is used to calcu-late the specific surface area as a function of the increasingtotal protein–pectin concentration (ESI, Table S1†). An increaseof the concentration of the emulsifier in the continuous phaseusually exhibits an increase in the specific surface area and adecrease in the oil droplet size and the polydispersity of the oildroplets.35 In addition, the finer droplet size with the increasein the total biopolymer concentration was also affected by theincrease in the viscosity of the dispersion (Table 2). It has beenreported that increasing the viscosity of the continuous phaseby using a protein-stabilizer results in smaller droplet sizesdue to the suppression of coalescence and increase in theviscous drag on the droplets.36

The wettability of the WPI–LMP particles by the liquidphases at each side of the interface can give an indicationabout the hydrophobic/hydrophilic characteristics of the par-ticles and the consequent performance of the particle-stabil-

Fig. 5 Droplet size distribution of oil droplets in fresh HIPE samplesstabilized by WPI (1.0%) and complexes (1.0 : 0.5%) formed at pH 3.5 (A),4.5 (B), and 5.5 (C), as well as a function of WPI : LMP concentration (%)in the complexes (formed at pH 4.5) (D).

Table 2 Apparent viscosity (at 10 s−1) and Z-average diameter of com-plexes and D[4,3] of HIPEs stabilized by complexes prepared at differenttotal concentrations of WPI : LMP at pH 4.5

WPI : LMP (%)Apparentviscosity (mPa s)

Z-Averagediameter (nm) D[4,3] (µm)

1.0 : 0.5 6 ± 1 255 ± 4 14.41 ± 0.591.5 : 0.75 16 ± 2 267 ± 5 12.32 ± 0.492.0 : 1.0 31 ± 4 302 ± 5 11.81 ± 0.512.5 : 1.25 45 ± 4 342 ± 6 10.31 ± 0.45

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ized emulsions. In the case of high internal phase oil-in-wateremulsions, it appears that optimal stabilization is obtained ifthe continuous phase wets the particle somewhat better thanthe dispersed phase does.37 In the current work, the wettingproperties of WPI–LMP particles were investigated by measur-ing the contact angle θw of a drop of water on a dry film ofWPI–LMP (0.5 : 0.25%) formed at pH 4.5 (Fig. S2, ESI†). Thevalue of θw (= 59.8 ± 1.0) revealed that the surface of the WPI–LMP complexes was partially wetted by water droplets. It indi-cates that the complexes are neither predominantly hydro-phobic nor hydrophilic, which is a requirement for Pickeringstabilization.37 In addition, the role of surface tensionreduction perhaps can also be associated with the surfaceproperties of the dispersion. The surface tension value forthe WPI–LMP dispersion (2.5 : 1.25%) was found to be47.7 ± 0.2 mN m−1 which is substantially lower than that ofpure water, 72 mN m−1. This result is comparable with aprevious report that the 2.5% β-lactoglobulin solution resultedin a surface tension of 50 mN m−1.38

Rheological properties

The rheological properties of HIPEs were evaluated to studythe influence of pH on the emulsion stability, and thus it wasconsidered to understand the microstructure-property links inthese complex emulsions. In food technology applicationparticularly, the rheological measurements can establish arelationship between the microstructure and the food texturalproperties since the data can be minimally interpreted to gainan insight into the structure breakdown of the food materialsduring mastication.39,40 The rheological measurements weredone on the HIPE samples on the following day after pre-paration, and were carried out on the samples stored at 5 °Covernight. The rheological properties of viscoelastic materialsare best determined through small-amplitude oscillatory shear(SAOS) rheometry, in which the parameters such as the visco-elastic limit (critical oscillatory stress), crossover point, andelastic (G′) and viscous (G″) moduli are identified.6

The data from amplitude sweeps conducted on HIPEsstabilized by WPI and complexes as a function of pH and con-centration are shown in Fig. 6A and 7A. For all of the samples,G′ was always greater than G″ at lower amplitudes, indicatingthat all the produced emulsions showed elastic or solid-likebehavior, but at higher amplitudes, a distinct crossover point(G″ > G′) was observed, suggesting the structural rearrange-ment or flow of the emulsion droplets at high applied stress.The stress values at the crossover point (or oscillatory yieldstress) for different emulsion samples are provided in the ESI(Tables S2 and S3†). The observed behaviour of these emul-sions is comparable to those of many other concentrated foodemulsions.41,42 Note that the measured G′ values for HIPEsstabilized by complexes were always higher than those ofHIPEs stabilized by WPI for all the pH values. This is likelyattributed to higher viscoelasticity of protein–polysaccharidelayers formed at the oil–water interface,14 besides benefitingfrom the high viscosity due to the close packing of oildroplets.

The aggregation of WPI and WPI : LMP particles directlyaffected the textural consistency of the emulsion samples.At pH 3.5, where the complexes were aggregated, the HIPEdisplayed a higher firmness than the ones stabilized by com-plexes formed at pH 4.5 and 5.5. In the case of emulsionsstabilized by WPI particles, soft gel-like textures were displayedat pH 3.5 and 5.5 in comparison to pH 4.5 where a lumpystructure with poor stabilization was obtained. Overall, thefirmest gel was produced by HIPEs stabilized by complexes atpH 3.5, a pH closer to the pI of the complexes. The controlledaggregation of complexes at pH 3.5 was associated with theaddition of pectin, which may cause the increase of particulate

Fig. 6 Amplitude stress sweeps (A), frequency sweeps (B), and flowmeasurements (C) carried out on HIPEs stabilized by WPI (1.0%) andcomplexes (1.0 : 0.5%).

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mass, thus leading to the HIPE with a firmer structure.However, this was not found in the HIPE stabilized by WPI atpH 4.5 in which the emulsion showed a lower G′ value causedby larger aggregation and poor stability (oiling-off ) in theabsence of LMP.

The increase in the total concentration of complexesresulted in a progressive increase in the gel strength as in-dicated by an increase in the values of the elastic modulus, G′,as well as an increase in the critical oscillatory stress (ESI,Table S4†). The increase in the solid dominant behavior withan increase in the total polymer concentration may be relatedto the stronger network formation in the continuous water

phase of the emulsion and as expected, a clear pattern ofincrease in the value was obtained with the increase in thetotal concentration.

The dependency of the material response to the appliedfrequency (i.e., the rate of deformation) was further studied bysubjecting the samples to a constant stress (2 Pa) that waswithin the region of linear response and varying frequencies(0.1 to 10 Hz). As seen from Fig. 6B and 7B, all HIPEs formedat different pH values and concentrations showed the charac-teristics of a strong gel (curves with slightly positive slopes),with higher values of G′ than G″ throughout the entirefrequency range. The inverse proportional relationship ofcomplex viscosity (η*) with increasing frequency (Hz) as a fun-ction of total biopolymer concentration also confirmed thestable gel structure of the samples (Fig. 6B). All HIPEs showeda very strong shear-thinning behaviour with almost no differ-ence in the steepness of the slope as clearly seen from the log–log plot shown in Fig. 6C and 7C. It is interesting to noticethat the viscosity of the sample with the weakest gel structure(i.e. HIPE prepared from the emulsion stabilized by 1.0 : 0.5%of complexes) is significantly higher than the viscosity of sun-flower oil (∼0.05 Pa s),43 even at the end of the flow measure-ment (at 20 s−1), hence confirming that no gel–sol transform-ation occurs under the applied stress probably due to the well-structured coverage of the oil–water interfaces by thecomplexes.

Stability of HIPEs

The stability of HIPEs in terms of phase separation wasmeasured by using an analytical photo-centrifuge, whichmeasures the transmittance of the emulsion during centrifu-gation. An increase in transmittance could indicate the separ-ation of a cream layer from the water phase, since no coalesc-ence was observed from the light transmission profiles(Fig. S3, ESI†). The creaming rate was determined from theslope of the curve of the integrated transmitted light intensityagainst time as explained in more detail in the Experimentalsection.44 Fig. 8A, B and Table S4 (ESI†) show the integraltransmission and creaming rate of HIPEs stabilized by WPIand complexes as a function of pH. All the emulsion sampleswithout complexation with LMP exhibited a greater degree ofcreaming when compared to emulsions stabilized by com-plexes, indicated by a dramatic increase of the transmittedlight signal in the initial 1000 s before reaching a plateau.As discussed earlier, a strong WPI : LMP complexationoccurred at pH 3.5 resulting in particles that couldn’t providebetter gravitational stabilization to emulsions as compared tocomplexes prepared at other pH values 4.5 and 5.5. On theother hand, HIPEs stabilized by WPI at all pH values werehighly susceptible to gravitational separation. The reason forthis can be explained by the fact that at pH values in the rangeof the isoelectric point of the adsorbed proteins, the decreasein electrostatic repulsion between the protein–protein mole-cules in the WPI solution would cause aggregation of protein-coated oil droplets.45 Moreover, the bulk phase viscosity of

Fig. 7 Amplitude stress sweeps (A), frequency sweeps (B), and flowmeasurements (C) carried out on HIPEs stabilized by complexes (formedat pH 4.5) as a function of concentration.

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WPI emulsions was comparatively much lower than that of theWPI : LMP emulsions.

The effect of total biopolymer concentration on the stabilityof the HIPEs stabilized by complexes is shown in Fig. 8C andTable S5 (ESI†). As the concentration increased, the creamingrate became lower. As shown in Fig. 8C, most stable emulsionscould be produced with a concentration of 1.5 : 0.75%(0.3 : 0.15% total weight in emulsion), as indicated by noremarkable increase of light transmission. The increase in theconcentration of complexes provides a better stabilizationsynergistic effect owing to a more efficient coverage of the oildroplets resulting in a smaller droplet size and also anincrease in the viscosity of the bulk phase.46

Since HIPEs stabilized by pH 4.5 and 5.5 complexes werevery stable towards coalescence during centrifugation, a versa-tile and powerful method has been proposed to inducecoalescence by ultracentrifugation. Ultracentrifugation was

performed at 25 °C, 20 000 rpm (70 952g) for 60 min. Afterultracentrifugation, phase separation of the HIPEs wasobserved. The appearance of HIPEs immediately after ultracen-trifugation is shown in Fig. 9A. In general, all emulsionsshowed three distinctive layers after ultracentrifugation, butthe emulsions stabilized by complexes formed at pH 4.5 and5.5 had a very small upper oil layer. Interestingly, after one dayof storage, the emulsions could be redispersed (Fig. 9B, shownby the red circle in the images).

In the current study, the result of oiling-off was in accord-ance with the creaming rate as shown by analytical photo-centrifugation data (Tables S4 and S5†). Oiling-off is generallyassociated with droplet aggregation and creaming. Thisphenomenon is attributed by droplet coalescence due to therupture of the interfacial film surrounding the oil droplets,thus leading to partial release of oil from the emulsion.47

In addition, electrostatic interaction may also modify theproperties of both protein and polysaccharide molecules,providing stronger mechanical properties against rupture.

Conclusion

Water continuous HIPEs containing an internal phase of 80 wt%(φoil = 0.82) with exceptional stability were successfully pre-pared by simply homogenizing aqueous dispersions of WPI–LMP complexes with sunflower oil in the absence of any low-molecular weight emulsifiers. We demonstrate that factors likepH conditions and total biopolymer concentration have astrong impact on the formation and stability of the resultantHIPEs. The rheological properties of the complex-stabilizedHIPEs can easily be tuned by varying the total concentration ofcomplexes at an appropriate pH of complexation. Our fabrica-tion process is straightforward and requires a relatively lowconcentration of biopolymers (only 0.2 g and 0.1 g of proteinand pectin used for 80 g oil stabilization). As opposed to manyother synthetic and non-food origin materials, non-covalentWPI–LMP complexes offer themselves as fully versatile ingredi-ents of natural origins. These advantages coupled with theirfavourable regulatory status could be of great interest in widerapplication domains such as for food and pharmaceuticalformulations.

Fig. 8 Integral transmission profiles by analytical photo-centrifugationof fresh HIPEs stabilized by WPI (1.0%) (A) and complexes (1.0 : 0.5%) (B)as a function of pH and as a function of concentration (at pH 4.5) (C).

Fig. 9 The images of HIPEs taken immediately after ultra-centrifugationat 70 952g (A) and one-day after ultra-centrifugation (B). The red circlesindicate that the bottom aqueous and upper oil layer could redispersedafter one-day storage.

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Acknowledgements

The first author would like to thank the BOF (Special ResearchFund) of Ghent University for providing financial support andMohd Dona Sintang for assisting in the cryo-SEM preparation.Ashok R. Patel would like to acknowledge the financialsupport from Marie Curie Career Integration Grant (callFP7-PEOPLE-2013-CIG, proposal no. 618157, acronymSAT-FAT-FREE) within the Seventh European CommunityFramework Programme. The Hercules foundation isacknowledged for its financial support in the acquisition ofthe scanning electron microscope JEOL JSM-7100F equippedwith a cryo-transfer system Quorum PP3000 T and OxfordInstruments Aztec EDS (grant number AUGE-09-029).

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