Stabilization of foams and emulsions by mixtures of surface active food-grade particles and proteins

Food Hydrocolloids 25 (2011) 627e638

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Stabilization of foams and emulsions by mixtures of surface active food-grade particles and proteins Brent S. Murray a, *, Kalpana Durga a, Anida Yusoff b, Simeon D. Stoyanov c a

Food Colloids Group, School of Food Science and Nutrition, University of Leeds, Leeds LS2 9JT, UK Faculty of Applied Sciences, Universiti Teknologi MARA, 40450 Shah Alam, Selangor, Malaysia c Unilever R&D Vlaardingen, 3133AT Vlaardingen, The Netherlands b

a r t i c l e i n f o

a b s t r a c t

Article history: Received 8 June 2010 Accepted 27 July 2010

Surface active biopolymers such as proteins can form films with particularly high interfacial elasticities and viscosities and these molecules are widely exploited as foaming and emulsifying agents in foods. Solid particles of the correct size and wetting characteristics can also be extremely effective stabilizers of foams and emulsions, although the underlying mechanism of stabilization is somewhat different. Relatively little is known about what happens when both surface active polymers and surface active particles are present together. This work presents recent findings on the effects of mixtures of proteins plus novel food-compatible surface active particles. The proteins include caseins and whey proteins. The surface active particles prepared include cellulose þ ethyl cellulose complexes, hydrophobically-modified starch granule particles and stable (non-spreading) protein-stabilized oil droplets. Interfacial shear rheology of adsorbed films was measured via a biconical bob apparatus and interfacial dilatational rheology was measured via a Langmuir trough type apparatus. The corresponding stability of bubbles to coalescence and disproportionation was assessed in separate experiments. Stability of oil-in-water emulsions was assessed via measurement of particle size distributions as function of time and visual assessment of the tendency to creaming and oiling off. In general, it is shown that the surface active particles on their own exhibit much lower measures of interfacial elasticity and viscosity than the proteins, but in combination with the proteins they appear to enhance the interfacial viscoelasticity considerably, with concomitant increases in bubble and emulsion droplet stability. There is little evidence of attractive interactions between the particles and the proteins, so a possible explanation of the increased stability is that the proteins increase the accumulation of particles at the interface, giving rise to increased jamming of particles at the interface. Ó 2010 Elsevier Ltd. All rights reserved.

Keywords: Pickering emulsions Proteins Foams Interfacial rheology

1. Introduction Emulsions and foams can be stabilized not only by surfactants, but also by solid particles. Emulsions of this type are termed Pickering emulsions, after the early work of Pickering (1907) although essentially the same phenomena were sketched out earlier by Ramsden (1903). Binks and Horozov (2006) have provided a survey of the subject of particle-stabilized colloids in general. The stabilization of food foams by particles has been reviewed by Murray and Ettelaie (2004) whilst Dickinson (2010) has more recently reviewed the stabilization of both emulsions and foams by particles in the context of foods.

* Corresponding author. Tel.: þ44 (0) 113 343 2962; fax: þ44 (0) 113 343 2982. E-mail address: [email protected] (B.S. Murray). 0268-005X/$ e see front matter Ó 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.foodhyd.2010.07.025

The key feature of particle-stabilized systems is that if the particles have the correct surface energy or contact angle with the interface, and also that they have a sufficiently large surface area, then the energy of desorption per particle can be of the order of several thousand kT. Such particles are thus effectively irreversibly adsorbed. This gives rise to significant advantages in that Ostwald ripening of emulsions, of even relatively soluble essential oils (Binks, Fletcher, & Holt, in press), plus disproportionation of bubbles of air in aqueous systems, can be almost completely arrested. Although proteins or low molecular weight surfactants (LMWS) can give excellent emulsification of oils in water and foamability of aqueous solutions, plus good stability to coalescence of the emulsions and foams produced, they are generally not good at preventing Ostwald ripening and disproportionation (Murray, 2007). One of the disadvantages of particles is that they may give relatively low foamability and rather coarse emulsions, principally because they are larger entities than molecules of surface active species. Mass

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transport and adsorption of particles to interfaces may therefore not be rapid enough to stabilize the smallest bubbles and droplets that can be formed during homogenization, mixing, whipping, etc. Since particles must have some hydrophobicity in order to adsorb (from the aqueous phase) this also means that they will also have a tendency to aggregate in the aqueous phase. This aggregation tends to slow down further the rate of coverage of the droplets or bubbles by the solid material. In this respect, surface active nanoparticles, i.e., particles with dimensions less than ca. 100 nm, that are colloidally stable by virtue of their small size may be an advantage. However, the deliberate addition or exploitation of some novel nanoparticles in foods may raise health concerns (Maynard et al., 2006). An advantage of particles compared to proteinaceous surfactants is that, particles can be made very hydrophobic and therefore dispersible in oil, where they can act as stabilizers of water-in-oil emulsions (Binks & Lumsdon, 2000). Indeed, natural fat crystals may act in this way in fatty spreads.(Rousseau, Ghosh, & Park, 2009) There are no natural oil-soluble proteins and a limited range of oilsoluble LMWS that can be used in foods to stabilize water-in-oil systems successfully. However, in this paper, we focus on systems where the continuous phase is aqueous, which constitutes the majority of food colloid products. Interesting questions arise if both surface active molecules and particles happen be present together, as will almost certainly be the case in most real food products. Any lipids or other LMWS present will probably adsorb to the particles and change their contact angle and adsorption characteristics. Proteins will adsorb to almost any type of surface, often with high affinity, and may therefore do the same. In addition, the interfacial coverage is likely to be dominated by the proteins or LMWS, since the particles will adsorb more slowly due to their slower mass transport. Whether proteins or LMWS might be able to displace any particles that just happen to adsorb to the bare interface first of all does not seem to have been thoroughly investigated. Work elsewhere (Binks, Kirkland, & Rodrigues, 2008; Cui, Yang, Cui, & Binks, 2010; Kostakis, Ettelaie, & Murray, 2007) has addressed competitive adsorption and interactions between model silica particles and cationic LMWS, and also between such silica particles and proteins (Kostakis et al., 2007). Perhaps what is even more relevant is how surface active particles might interact with an adsorbed protein film that has already been formed e the particles might be repelled from the film, become incorporated in it, or might even displace the protein from the interface. Dickinson (2010) concludes his review of studies of particlestabilized colloids with an observation that, despite the large amount of research particle-stabilized systems, relatively little of this research is directly applicable to foods because the particles that have been used are not permissible in foods. There are some notable exceptions such as: wax crystals (Campbell, Holt, Stoyanov, & Paunov, 2008, Campbell, Stoyanov, & Paunov, 2009a, 2009b; Binks & Rocher, 2009); ethyl cellulose (Campbell, Stoyanov, & Paunov, 2009a, 2009b); modified CaCO3 (Zhou, Cao, Liu, & Stoyanov, 2009); thermally induced protein aggregates (Paunov et al., 2007; Unterhaslberger, Schmitt, Shojaei-Rami, & Sanchez, 2007); proteinepolysaccharide complexes (Schmitt, Aberkane, & Sanchez, 2009; Turgeon, Schmitt, & Sanchez, 2007), or protein/ polysaccharide coacervates (Moschakis, Murray, & Biliaderis, 2010; Schmitt et al., 2005). However, aggregates, complexes and coacervates are obviously particles that are complex in terms of their structure and composition. In this work we describe three types of particle of fairly simple structure and composition that are definitely compatible with foods and how these interact with proteins in foams and an oil-in-water emulsion. The particles are: (i) stable, non-spreading oil droplets, (ii) hydrophobically-modified cellulose fragments and (iii) hydrophobically-modified starch granule particulates.

2. Materials and methods 2.1. Materials Gelose 80 starch was obtained from Penford Food Ingredients, USA. Octenyl Succinic Anhydride (OSA) was donated by the Dixie Chemical Company, Pasadena, Texas, USA. Sodium azide, potassium dihydrogen phosphate, disodium phosphate, hydrochloric acid, acetone, sodium thiocyanate, urea and sodium hydroxide were of AnalR grade and obtained from SigmaeAldrich (Gillingham, UK). Ethyl cellulose (EC) (product code 02366, 48.0e49.5% ethoxyl content), n-tetradecane (99%), glucono-d-lactone (GDL) (99%), bovine b-lactoglobulin (BL) (three times crystallized, lyophilized, desiccated, lot no. 21K7079, containing variants A and B), Nile Blue and Nile Red were also obtained from SigmaeAldrich. Spray-dried sodium caseinate (SC) (>82 wt.% dry protein, <6 wt.% moisture, <6 wt.% fat and ash, 0.05 wt.% calcium) was supplied by DMV International (Veghel, Netherlands). Commercial whey protein isolate (BiPro) was obtained from Davisco Foods (MN, USA) and contained 97.7% protein, 0.3% fat, 1.9% ash and 4.8% moisture. Water purified by treatment with a Milli-Q apparatus (Millipore, Bedford, UK), with a resistivity not less than 18.2 MU cm, was used for the preparation of all solutions. Aqueous solutions of SC and BL were prepared by dispersing the required amount of protein in buffered Milli-Q water containing 0.02% w/v sodium azide under gentle stirring for 4 h at room temperature. The buffers consisted of 0.05 mol dm3 phosphate þ 0.05 mol dm3 NaCl, at pH 6 or pH 7. 2.2. Preparation and characterization of oil-in-water emulsions Two types of emulsion were prepared. One type was a 30 vol.% oil-in-water (O/W) emulsion formed from 1 wt.% protein solution and n-tetradecane via a Shields S-500 high-pressure homogenizer. This type of emulsion was added as a type of ‘droplet particle’ to foams to see how this influenced bubble stability. The same protein solution was used to stabilize the emulsion as was used to stabilize the bubbles. The second type of emulsion was a 20 vol.% O/W emulsion formed from a 3 wt.% aqueous dispersion of the surface active starch particles and n-tetradecane. These starch-stabilized emulsions were prepared using a high-pressure jet homogenizer (Burgaud, Dickinson, & Nelson, 1990) operating at 300 bar. Emulsions were stored in sealed glass test tubes (height 75 mm, diameter 24 mm) immediately after preparation at room temperature. Emulsion droplet size distributions were measured by static multi-angle light scattering via a Mastersizer Hydro 2000 (Malvern Instruments, Malvern, UK). The refractive indices of aqueous phases and n-tetradecane were taken as 1.330 and 1.429, respectively. Average droplet sizes were characterized in terms of the Sauter mean diameter d32 or volume mean diameter d43 defined by:

P i

dab ¼ P i

ni dai ni dbi

(1)

where ni is the number of the droplets of diameter di. All measurements were made at room temperature on at least two freshly prepared samples. The size of the protein-stabilized n-tetradecane droplet particles was d43 ¼ 0.59  0.05 mm. The droplet sizes of the starch-stabilized emulsions are discussed below. 2.3. Preparation of surface active cellulose particles A range of surface active cellulose particles have been produced using a variety of techniques and this work is described in detail elsewhere (Murray, Durga, de Groot, & Stoyanov, in preparation).

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In this paper just one type of cellulose particle was used. Briefly, this was prepared as follows. Tencel (1.7dtex 3 mm bright Nonwovens H400431) was used as the source of cellulose, obtained from Lenzing fibres Ltd., Derby. Tencel itself is a pure form of cellulose obtained from wood pulp by direct dissolution in N-methylmorpholineN-oxide. The initial mean length of the Tencel fibres as received was 3 mm, with a mean width of 20 mm. Plant or bacterial cellulose can be made hydrophobic by complex formation with suitable additives (Andresen & Stenius, 2007; Wege, Kim, Paunov, Zhong, & Velev, 2008). However, in order to act as an efficient stabilizer of acceptably small bubbles or emulsion droplets the cellulose particle (fibre) size must also be suitably reduced. Physical, chemical and enzymatic methods of breaking down Tencel have been attempted, but physical breakdown appeared to be the most efficient in terms of quickly and easily producing fairly large quantities of material. The Tencel was hammer-milled four times for 2 min in a C & N eight inch laboratory hammer mill (Christy Turner Ltd., Ipswich), with 1e2 min between each milling and then cryogenically milled using a SPEX CertiPrep 6750 cryogenic freezer mill (Metuchen, USA). In the freezer mill the sample was pre-cooled for 12 min and then processed for 3 cycles. Each cycle was 2.4 min long and there was a time interval of 2 min between each successive cycle. This had the effect of reducing both the mean fibre length and also fraying the Tencel particles to release thinner fragments. Most of the fragments ranged in width from 5 to 20 mm and in length from 0.5 to 70 mm, with the mean the aspect ratio of the fragments between 3 and 5, although there were smaller fragments that were hard assess in terms of their size and shape. It is seen that the final aspect ratio was much lower than that before milling. Smaller fragments can be produced by more extensive physical and chemical procedures, but these were not used in the work reported here. The method used to make the cellulose particle surface active was similar to that described by Campbell et al. (2008) in order to produce surface active shellac fibres. The cellulose particles were dispersed in an organic solvent (acetone) at 50  C and an equal weight of ethyl cellulose (EC) added. The dispersion was mixed via a magnetic stirrer whilst an equal volume of pH 6 aqueous buffer was added. EC is insoluble in water and precipitates onto the cellulose as the dilution takes place, giving rise to hydrophobic patches on the particles that make them surface active. The resultant EC-cellulose particles are hereafter referred to as ‘cellulose complexes’. Different ratios of EC to cellulose have been experimented with (Murray et al., in preparation), as well as different overall concentrations of cellulose and EC during the dilution procedure. However, the basic findings are that if too high an EC : cellulose ratio is used, then the particles become too hydrophobic and aggregate together to form a viscous ‘gel’ that is unsuitable for further use. If too low a ratio is used then the cellulose particles are not surface active enough to adsorb to bubble surfaces. The complexes used here were initially formed at concentrations of 1 g of Tencel þ 1 g of EC per 100 ml of solvent, before allowing the solvent to evaporate. The dispersion of complexes was diluted with the appropriate volume of buffer for subsequent experiments. The exact nature of the interactions between the EC and the cellulose is not clear. It is assumed that hydrogen bonding takes place between the polysaccharide chains of the EC and the cellulose, although the EC appears to be localised as distinct globules along cellulose fibres (results not shown). What is clear is that once complex formation has taken place, the EC seems to be irreversibly adsorbed to the cellulose. EC dispersed on its own via the same procedure gave reasonable foamability, but very poor foam stability and was not studied further. 2.4. Preparation of surface active starch particulates The details of the procedures used to generate various surface active starch particulates have been described in detail elsewhere

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(Yusoff & Murray, 2011). In this study only one type of modified starch was used and a summary of the preparation details is as follows. The size of the native Gelose 80 starch granule material was reduced by cryogenic milling as for the Tencel, resulting in particles in the size range 0.5e15 mm. The starch was made hydrophobic by treatment with octenyl succinic anhydride (OSA) according to the procedures described by Bhosale and Singhal (2006). The pH of a slurry of approximately 20 wt.% starch in Milli-Q water was adjusted to 8.0  0.2 by addition of 2% NaOH solution. To this mixture, OSA at 3%, based on the weight of starch, was added drop-wise for 2 h, maintaining the pH at 8.0  0.2 by the addition of NaOH. The reaction mixture was left for 24 h at 30  C then the pH of the mixture was adjusted to 6.5 by addition of 2% v/v HCl. The resulting slurry was washed with acetone (1) and water (3) and then filtered through a similarly washed Whatman No. 1 filter paper. The slurry was scraped from the filter paper and redispersed in pH 7 buffer with gentle stirring. The degree of substitution of the starch was determined using the method of Whistler and Paschall (1967) and measured as 0.03  0.005. Before use in experiments, the starch slurries were diluted to the appropriate concentration with the same buffer and mixed for 30 s at 24 000 rpm with T25 S7 Ultraturrax high shear mixer (Janke & Kunkel, Staufen, Germany) to ensure as complete dispersion of the starch as possible. Some of the starch dispersions were also passed through the jet homogenizer on their own (i.e., with no oil phase present) at 300 bar before using them to stabilize emulsions. 2.5. Confocal microscopy A Leica TCS SP2 confocal laser scanning microscope (CLSM), mounted on a Leica Model DM RXE microscope base, was operated in fluorescence mode. Nile Blue was used to stain the starch particles and Nile Red was used to stain the oil phase, as described elsewhere (Yusoff & Murray, 2011). Approximately 80 ml of the stained sample was placed into a laboratory-made welled slide, (Moschakis, Murray, & Dickinson, 2005) filling it completely. A cover slip (0.17 mm thickness) was placed on top of the well, ensuring that there was no air gap (or bubbles) trapped between the sample and cover slip. The samples were scanned at 24  C, using 10 (NA 0.3) or 40 (NA 1.25) oil-immersion objective lenses, approximately 10e20 mm below the level of the cover slip. Images were recorded at a resolution of 1024  1024 pixels and processed using the image analysis software Image J. 2.6. Measurement of stability of bubbles and foams to coalescence and disproportionation The bubble coalescence cells and their operation have been described in detail previously (Murray, Cox, Dickinson, Nelson, & Wang, 2007; Murray, Dickinson, Lau, & Schmidt, 2005; Murray, Dickinson, & Wang, 2009a) and only brief information is given here. Bubbles are created in specially designed cells under conditions such that the majority of bubbles are stable to coalescence until instability is deliberately induced by suddenly lowering the pressure, whence the proportion of induced instability is quantified. In one type of cell coalescence of a single layer of bubbles with a planar airewater (AeW) interface is measured within a flexible square barrier positioned in the interface. Bubbles with a typical size range of 100e300 mm diameter are injected beneath the planar interface within 1e2 min, whence they rise to the interface. The barrier is then expanded and the air pressure above the interface simultaneously decreased in order to achieve simultaneous and equal expansion of the bubbles and the planar interface. The bubbles are observed via a microscope and the number fraction of bubbles that are stable to coalescence on expansion, Fs, is

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determined from the digitally recorded images. Experiments were repeated at least 3 times and results based on the mean values of Fs are reported. The minimum reproducibility in these mean Fs values was 0.05. The measurement of the stability of foams to coalescence is based on the same principle of applying a pressure drop in order to induce instability. This method has also been described in detail elsewhere (Murray et al., 2007). In these experiments the aqueous phase was carefully added to the cell to avoiding foaming and then a foam was formed in a controlled way by injecting air at a controlled flow rate beneath the surface of the solution, via a 100 ml syringe attached to a long thin capillary. The cell was sealed and the foam subjected to a similar pressure decrease as in the single bubble layer experiments. The bubbles in the threedimensional foam expand against each other and the fraction of bubbles stable to coalescence (Fs) is estimated by image analysis of the layer of foam visible near the wall of the cell. Experiments were repeated at least 3 times and results based on the mean values of Fs are reported. The minimum reproducibility in these mean Fs values for the foams was 0.06. The agreement between the single bubble layer and foam bubble stability measurements is generally good, except under conditions where it is difficult to produce a foam possessing fairly uniform sized bubbles in the first place, i.e., before applying the pressure drop (Murray et al., 2007). In the measurement of bubble shrinkage, i.e., disproportionation, bubbles are injected beneath the AeW interface in the same type of cell as used for the measurement of coalescence in the single layer of bubbles. After bubble injection the bubble size is simply monitored as a function of time. The bubbles slowly shrink due to the dissolution of gas into the aqueous phase and through the planar interface. The process has been modelled in detail (Dickinson, Ettelaie, Murray, & Du, 2002; Ettelaie, Dickinson, Du, & Murray, 2003). For convenience here, we have simply reported the time, ts, for individual bubbles of radius 150 mm to dissolve ‘completely’ e meaning that they were no longer clearly visible because their size was less the optical resolution of the microscope (ca. 3 mm). Experiments were repeated at least 2 times, starting with bubbles that had the same starting radius 5 mm and results based on the mean values of ts are reported. The minimum reproducibility in these mean ts values was 10 min.

2003). Experiments were repeated at least 3 times and results based on the mean values of hi are reported. The minimum reproducibility in these mean hi values was 20%. A Langmuir trough apparatus with a flexible square rubber barrier, as described by Murray, Xu and Dickinson (2009b), was used to measure the dilatational rheology of the adsorbed films. The inside dimensions of the trough in this case were 14.3 cm2 and the rubber barrier was able to expand from 4.5 to 12 cm2. After filling the trough to the required level with the aqueous phase, the surface was aspirated away via a clean Pasteur pipette and vacuum pump. The interface was then left for 2 min before suddenly expanding the interface and recording the change in surface tension (Dg) with time. The 2 min waiting time was to match, as far as possible, the time taken to inject bubbles beneath the planar interface in the bubble coalescence cell. The planar interface in the trough was subjected to the same rate of area strain dln A/dt and the same relative increase in area, A/A0 (where A0 and A are the initial and final areas, respectively) as in the bubble coalescence measurements. Further details are given elsewhere (Murray, Dickinson, & Wang, 2009a; Xu, Dickinson, & Murray, 2008). As in our previous work (Murray et al., 2009b; Xu, Dickinson, & Murray, 2007, 2008), it was found that the initial gradient of the Dg versus ln A curve was a convenient measure of the dilatational rheological response of the films. The Dg versus ln A data over the first 5% of the total time of the expansion period were fitted to a straight line and the gradient of this line termed the ‘average initial dilatational elasticity’, 3*init. This parameter is actually a complex modulus, although more detailed analysis shows that the elastic (storage) component generally dominates under these conditions of expansion. The fit to a straight line was always good enough to give a regression coefficient > 0.97. A further justification for analyzing the data in this way is that, when bubbles coalesce in the pressure drop tests, they tend to do so during the first few % the overall expansion. Experiments were repeated at least 3 times and results based on the mean values of 3*init are reported. The minimum reproducibility in these mean 3*init values was 20%.

3. Results and discussion 3.1. Oil droplets as stabilizing particles

2.7. Measurements of interfacial shear viscosity (hi) and complex dilatational elasticity (3*init) To measure the interfacial shear viscosity of the adsorbed films, a two dimensional Couette-type interfacial viscometer (Murray, 2002), was operated in a constant shear-rate mode, as described previously (Murray et al., 2009a). A stainless steel biconical disc (radius 14.5 mm) is suspended from a thin torsion wire with its edge in the plane of the AeW or oilewater (OeW) interface of the solution contained within a cylindrical glass dish (radius 72.5 mm). The constant shear-rate apparent interfacial viscosity, hi, is given by the equation:

hi ¼

gf

u

Kðq  q0 Þ

(2)

where K is the torsion constant of the wire; q is the equilibrium deflection of the disc in the presence of the film; q0 is the equilibrium deflection in the absence of the film, i.e., due to the bulk drag of the subphase on the disk; gf is the geometric factor and u is the angular velocity of the dish. Torsion wires of varying thickness and lengths were use to achieve the appropriate torsion constant. A fixed value of u ¼ 1.27  103 rad s1 was employed throughout, for comparison with measurements elsewhere (Borbas, Murray, & Kiss,

Fig. 1 illustrates the significant increase in stability to coalescence of air bubbles (measured in the single bubble layer experiments) when a low volume fraction (0.25%) of n-tetradecane droplets is included. The bubbles were formed in solution to which the droplets had already been added and it should be emphasized again that these droplets behave as completely stable entities e there

4 3 RFs 2 1

5

pH

6

7

Fig. 1. Ratio of the number fraction of air bubbles stable to coalescence in the presence of 0.25 vol.% oil droplets to the number fraction stable in the absence of oil droplets (RFs) versus pH for: 1 wt.% b-lactoglobulin (6); 1 wt.% sodium caseinate (C).

B.S. Murray et al. / Food Hydrocolloids 25 (2011) 627e638

is no evidence for them entering or spreading at the AeW interface (as in the case of studies under very different conditions e for example see Hotrum, van Vliet, Stuart, & van Aken, 2002). The relative stability (RFs) has been plotted as function pH where RFs equals the ratio of Fs in the presence of the droplets to Fs in the absence of the droplets. Values of RFs > 1 therefore mean that the system is more stable when droplets are present, which is clearly true for both BL and SC across the pH range 4.5e7. The strong increase in RFs for BL as the pH is lowered is most likely due to the decrease in the net charge on the adsorbed film of BL on the bubbles and the droplets, the isoelectric pH of BL being ca. pH 5.3. Although this will tend to reduce repulsion between interfaces, which might be expected to decrease stability to coalescence, it also tends to give rise to enhanced protein adsorption and higher values of the interfacial shear viscoelasticity (Murray, 2002) as mutual repulsion between adsorbed protein molecules is decreased. It may also lead to increased co-adsorption of emulsion droplets to the protein film on the bubbles, as indicated in the schematic diagram in Fig. 2, although as yet we have no direct evidence for this. As the pH is lowered in the SC system RFs seems to follow the same sort of trend, but below about pH 5.8 there is a sharp reversal in RFs, so that by about pH 5.5 there is little enhancement in coalescence stability with droplets. The reason for this is that, below pH z 5.8 SC begins to precipitate out of solution and the bubbles both with and without droplets become very unstable. In contrast, BL remains soluble as the pH crosses the pI of this protein. In previous work (Murray et al., 2009a) we have described how stability to disproportionation is also increased by the presence of the droplets, although the increases are not as marked as with the coalescence stability. It has also been shown that the increase in stability in the SC system correlates quite well with a marked increase in the interfacial shear viscosity in the presence of the droplets. This is not simply due to creaming of the n-tetradecane droplets to the interface, since the same effects were also observed with neutrally buoyant 1-bromohexadecance droplets of very similar size. Thus, as Fig. 2 implies, there seems to some sort of positive association of the droplets with the adsorbed protein film at the airewater interface, even at neutral pH when there should be a significant electrostatic repulsion between the interfaces. On the other hand, the conformation of the protein at the two types of interface is not necessarily the same (Murray, Færgemand, Trotereau, & Ventura, 1998; Pradines, Kragel, Fainerman, & Miller, 2009) and possibly this could give rise to some sort of weak attraction between the AeW interface and the OeW interface.

Fig. 2. Schematic diagram indicating the possible mechanism by which proteinstabilized oil droplets influence the strength of an adsorbed film of the same protein at the airewater interface as the pH is decreased towards to the pI of the protein.

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Emulsion droplets may therefore be used as a sort of ‘particle’ to strengthen the adsorbed protein film and give enhanced stability. Of course the interior of the particles is not solid but completely liquid. Many other real systems, e.g., ice cream mixes and dairy toppings contain such mixtures of bubbles and droplets, as well as other particles, such as intact casein micelles that may reach into the same size range and possibly act in similar ways, although this has not yet been studied. The droplet particles have the advantage that the particles are relatively well-defined in terms of their size, shape and surface properties. However, since they are unlikely to adsorb to the bare interface and remain there as stable entities, it can be argued that they are not acting as true particle stabilizers of the bubbles. It is possible to produce solidified droplets by emulsifying higher melting point oils above their melting point and then cooling them well below their melting point, which can then be studied for their effects on whipping and bubble stability (Allen, Dickinson, & Murray, 2008; Allen, Murray, & Dickinson, 2008). Even so, the droplets themselves still require a stabilizing layer of protein or other surfactant in the first place, which complicates the interpretation of their mode of action. For this reason, we now turn to the cellulose complexes as foam stabilizers, which are truly solid, surface active particles in their own right. In addition, it is known from both theoretical and experimental studies (Binks & Horozov, 2006) that solid particles of a higher aspect ratio (i.e., not spherical) have clear advantages in terms of foam stabilization. 3.2. Cellulose complexes as stabilizing particles Simply shaking the suspension of cellulose complexes in sealed test tubes revealed that they were capable of forming foams that were highly stable. In most cases some smaller bubbles initially tended to disappear, whilst other bubbles seemed to become slightly larger in the first 1e2 h of storage, but after this time all the bubbles remained stable for many weeks, indicating remarkable stability to disproportionation. Fig. 3 shows typical CLSM images of the particle-stabilized bubbles, where Nile Blue has been used to highlight the complexes, which appear as the bright regions in this gray scale image. One of the images (Fig. 3(d)) shows bubbles after they have been stored for 24 h and distinctly non-spherical ‘bubbles’ can be seen, indicative of a highly compressed and buckled interface that is resistant to bubble shrinkage via gas dissolution (i.e., disproportionation). Although the air bubbles were highly stable to coalescence and disproportionation once they had been formed, the cellulose complex dispersions did not show high foamability and it was difficult to produce many very fine bubbles, i.e., less than ca. 0.5 mm diameter. Undoubtedly this was due to the large size of the complexes and their aggregates which, as has already been noted, will restrict the speed of their coverage of the interface. It was therefore of interest to mix the cellulose complex particles with a protein as a better foaming agent and see what foam properties were obtained. Fig. 4 presents coalescence data for foams formed from mixtures of cellulose complexes þ whey protein isolate (WPI) at pH 6. The relative stability to coalescence RFs, where RFs is the ratio of Fs in the presence of the complexes to Fs in the absence of the complexes (i.e., in the same manner as in Fig. 1 for the droplet particles) has been plotted as a function of the bulk concentration of WPI. The cellulose complex concentration was fixed at 0.1 wt.%. It is seen that at low WPI concentrations there is a significantly higher stability to coalescence in the presence of the complexes. As the WPI concentration is increased stability decreases and this is probably due to the higher surface activity of the WPI so that, by ca. 0.5 wt.% WPI the AeW interface and bubble stability is dominated by the protein. At very low WPI concentrations alone stable bubbles cannot be produced at all. For reference, the absolute Fs for WPI

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Fig. 3. Examples of confocal microscopy images of air bubbles freshly stabilized by 1 wt.% of the surface active cellulose complexes at pH 6 ((a)e(c)). The complexes were stained with Nile Blue and appear as bright regions. Fig. 3(d) shows a foam that has been aged for at least 24 h, highlighting the appearance of non-spherical bubbles.

alone at the lowest protein concentration (0.01 wt.%) depicted in Fig. 4 was Fs ¼ 0.07. Fig. 5 shows that there was a similar improvement in stability to disproportionation when adding 0.1 wt.% complexes to WPI, BL and also SC. The relative stability Rts, the ratio of time to shrink in the presence of the particles to the shrinkage time in the absence of the particles, has been plotted as a function of bulk protein concentration, Cp. The behaviour of the WPI and BL systems is similar in that the greatest enhancement in stability appears to occur at low Cp, for probably the same reasons as with the coalescence stability, in that protein probably dominates the interface at short times and particularly at higher Cp. Pure protein films are unable to provide resistance to bubble shrinkage (Dickinson et al., 2002), unlike adsorbed films of solid particles. In contrast, the stability with the SC þ cellulose complexes seems to continually increase as the SC

concentration increases. The reasons for this are not clear e possibly a higher SC concentration is required to ensure some incorporation of complexes into the film and produce increased in stability. Fig. 6 shows measurements of the complex dilatational elasticity 3*init of the proteins in the presence and absence of the cellulose complexes at the same compositions as shown in Figs. 4 and 5. It is seen that across most of the compositions the inclusion of the complexes causes only a slight change (increase) in 3*init. The only exception is WPI where there is a more significant increase in 3*init between Cp ¼ 0.01 wt.% and 0.1 wt.% in the presence of the complexes, followed by a sharp decrease to similar values without complexes by Cp  0.5 wt.%. The measurement 3*init of relates to the first fraction of the fairly rapid expansion of the interface and as such one would expect the 3*init parameter to be more relevant to the coalescence measurements induced by the pressure drop rather

4 5

3 RFs

Rts 3

2 1

0.0

0.2 0.4 wt% WPI

0.6

Fig. 4. Ratio of the number fraction of air bubbles stable to coalescence in the presence of 0.1 wt.% cellulose complexes to the number fraction stable in the absence of complexes (RFs) versus wt.% whey protein isolate at pH 6. (The line drawn through the points is to guide the eye.)

1

0.0

0.5 Cp/ wt.%

1.0

Fig. 5. Ratio of the time of bubble shrinkage in the presence of 0.1 wt.% cellulose complexes to the time of bubble shrinkage in the absence of complexes (Rts) as function of bulk protein concentration (Cp) at pH 6 for: whey protein isolate (7); b-lactoglobulin (B); sodium caseinate (>).

B.S. Murray et al. / Food Hydrocolloids 25 (2011) 627e638

60 ε*init / mN m 30

-1

0 0.0

0.5 Cp/ wt.%

1.0

Fig. 6. Complex dilatational modulus 3*init at the AeW interface at pH 6 in the presence of 0.1 wt.% cellulose complexes (open symbols) and in the absence of cellulose complexes (filled symbols) versus bulk protein concentration (Cp) for whey protein isolate (7, ;); b-lactoglobulin (B, C); sodium caseinate (>, A).

than resistance to disproportionation, since the latter involves a much slower change in interfacial area and interfacial compression rather than expansion. Comparison of the coalescence stability measurements in Fig. 4 with the dilatational viscoelasticity measurements in Fig. 6 for the WPI þ complexes does not suggest a simple the relationship between Fs and 3*init. On the other hand, this might be expected, since there are a large number of competing processes that might take place on expanding the interface in the presence of the mixture of surface active proteins and cellulose particles. Besides interfacial re-organization of the mixed proteineparticle film on expansion, this will be followed by adsorption of protein, complexes, or protein þ complexes, where the protein has adsorbed onto the complexes. Furthermore, these three types of species may compete for adsorption sites, but their rate of diffusion to the interface will depend on the equilibrium concentration of the same species in the subphase and how a shift in this concentration affects the aggregation of cellulose complexes with each other. How all this affects the transient change in g on expansion is hard to predict. However, similar measurements on the system of SC þ oil droplet particles as a function of pH (Murray et al., 2009a) suggest a stronger correlation of stability to coalescence with the interfacial shear viscosity, hi. For this reason hi was also measured for some of the protein þ cellulose complex mixtures. Fig. 7 shows hi at the AeW interface for 0.1 wt.% protein þ 0.1 wt.% complexes. A number of other compositions have been studied and will be reported elsewhere (Murray et al., in preparation), but this combination of concentrations has been the focus of these studies because it was generally the combination that gave particularly high stability to both coalescence and disproportionation, as seen in Figs. 4 and 5. The measurements were generally not continued much beyond 3e4 h, since this was the

ηi /

4

N s m-1 2 0

0

1

t/ h

2

Fig. 7. Interfacial shear viscosity (hi) versus time (t) at the AeW interface at pH 6 in the presence of 0.1 wt.% cellulose complexes (open symbols) and absence of cellulose complexes (filled symbols) for 0.1 wt.% whey protein isolate (7, ;); b-lactoglobulin (B, C); and sodium caseinate (>). The values for sodium caseinate alone and the fibres alone were barely measurable, as indicated by the dashed line.

633

usual time-scale of the stability measurements. The complexes on their own gave negligible hi. The complexes do tend to settle out under the conditions of the measurement of hi, which involves little agitation compared to the conditions of preparation of foams, etc. However, in some experiments vigorous stirring of the subphase was intermittently applied but this did not result in any significant increase in the values of hi. In contrast to the values for the complexes alone, hi values for WPI and BL were easily measurable and very much higher. WPI gave lower values than for the (pure) BL, but this is probably due to the presence of impurities in the WPI that affect the protein cross-linking at the interface. Most significant, however, was the considerable large increase in hi when complexes were present along with the proteins. This was most marked for SC þ complexes, which gave values of hi 2e3 orders of magnitude higher than SC on its own. (On its own 0.1 wt.% SC gave negligible values of hi, like the complexes on their own.) In fact, after ca. 3 h adsorption, the SC þ complex system gave similar values to the BL þ complex system. Globular proteins like BL generally give much higher values of hi than the individual caseins or caseinate, due to the greater tendency for unfolding and crosslinking of globular proteins at the interface (Murray, 2002, 2010). The large increase in hi when the complexes are added to the proteins suggests some sort of attractive interaction between the two components that leads to overall strengthening of the adsorbed film. This increase in hi correlates with the general increase in stability to coalescence and disproportionation when the complexes are present, as was also observed for the case of oil droplets as particles. The types of interaction responsible might include hydrogen bonding between the cellulose and the proteins, attractive hydrophobic interactions between the hydrophobic EC regions of the complexes or even electrostatic interactions between the complexes and the proteins e although the complexes formed by the physical methods described do not show much evidence of surface charge. [Cellulose degraded via the common method of treatment with concentrated sulfuric acid can develop significant negative charge due to the grafting of sulfate groups to the cellulose (Marchessault, Morehead, & Walter, 1959).] On the other hand, it may be that the particles simply act as an ‘inactive’ filler, in that they are trapped in the adsorbed protein layer though their adsorption to the interface, but not particularly bonded to the protein network. As such, they may still provide increased resistance to deformation of the interface and aid bubble stability. In order to test for hydrogen bonding interactions between the complexes and the proteins the effect on hi of changing the temperature of the films was measured. Films were heated up to 40  C, cooled down to 15  C and then heated back up to 25  C, the original temperature of measurement. This was done starting with films that were 24 h old, so that hi was not significantly changing with time. The films were held at each new temperature for 30e40 min before making the measurement of hi. Over this temperature range, if hydrogen bonding is responsible for the attractive interactions between the film components and the film strength, there should be a significant decrease in hi between 15 and 40  C. On the other hand, the temperature of 40  C is not high enough to cause irreversible thermal denaturation of WPI, BL or SC. Fig. 8 shows an example of the results obtained for 0.1 wt.% WPI and 0.1 wt.% WPI þ 0.1 wt.% complexes. The arrows indicate the direction of the temperature change. It is seen that there is a significant decrease in hi between 15 and 40  C. The change is apparently completely reversible, since hi measured at the beginning of the cycle (at 25  C) was almost identical to the value measured at the end of the cycle (also at 25  C). Whilst the decrease in hi with temperature may point to hydrogen bonds as being important to the strength of the WPI films, it seen that the gradient of hi versus temperature plot for WPI þ complexes is almost the same as for WPI

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B.S. Murray et al. / Food Hydrocolloids 25 (2011) 627e638

3 ηi /

-1

Nsm 2

1

10

20

o

T/ C

30

40

Fig. 8. Interfacial shear viscosity (hi) versus temperature (T) after 24 h adsorption at the airewater interface at pH 6 for: 0.1 wt.% whey protein isolate (6); 0.1 wt.% whey protein isolate þ 0.1 wt.% cellulose complexes (7). The arrows indicate the direction of the temperature change, i.e., 25 to 40 to 15 and back to 25  C.

alone. If the increase in temperature disrupted hydrogen bonding between the WPI and the complexes then one might expect a transition from the data set for the mixture to that of the WPI alone e but this was not observed. The same data from Fig. 8 have been plotted in a different way in Fig. 9, along with corresponding data for the BL systems and also some old data for gelatin and a different sample of SC measured under slightly different conditions (Castle, Dickinson, Murray, & Stainsby, 1987; Murray, 1987). In Fig. 9 hi values are scaled relative to the values at 25  C and the logarithm of these scaled values is plotted against reciprocal absolute temperature (T). Plotting the scaled values allows one to compare more easily data covering different ranges of absolute hi. For example, the absolute values of hi for SC are always very much lower than the values observed for BL. The reason for plotting the logarithm of hi versus (1/T) is to test for conformity to the MooreeEyring theory (Moore & Eyring, 1938) of surface viscosity. The MooreeEyring theory predicts a linear dependence of ln(hi) versus on (1/T) based on a simple activation energy barrier to the flow of units within the interface. Formally, it may be stated as:

hi ¼

  DG þ pDA h exp DA kT

(3)

where h ¼ Planck’s constant; DA ¼ area of kinetic flow unit; p ¼ surface pressure and DG ¼ activation energy. Making the

η

η

reasonable assumption that p does not significantly vary over the temperature range for the fully saturated films examined here, and also that DA is temperature independent, hi should vary linearly with (1/T). The gradient of the plot can be used to estimate DG. The assumption of invariant DA is difficult to prove and DA is difficult to access independently for adsorbed films. From spread protein monolayer studies MacRitchie (1986) has estimated that DA represents sections of polypeptide 6e8 amino acid residues long for most proteins. Irrespective of doubts over their theoretical interpretation, the data in Fig. 9 indicate a number of features. Firstly, the temperature dependence of SC on its own is very much greater than that of the pure globular protein BL (or the less pure WPI). In fact the SC data seem to obey the MooreeEyring theory quite well. It should be emphasized again that all the data in Fig. 9 represent completely reversible changes in hi when the temperature is changed. The temperature dependence of gelatin is also much higher than for BL (or WPI) but the data falls into two distinct regions, above and below approximately 30  C (303 K). Above 30  C the temperature dependence is similar to that for SC, but below 30  C the temperature dependence is less than for SC, but still significantly greater than for BL. The different temperature dependencies are summarized in Fig. 10, where DG obtained by fitting a straight line to the data below 30  C has been plotted against the absolute values of hi at 25  C after 24 h. It is interesting that the lower the value of hi, the greater DG, i.e., the weaker films are more susceptible to weakening by an increase in temperature. The origins of these differences for the proteins on their own deserve further study. For example, the results may provide evidence for the globular protein film existing in a glassy state (Cicuta, Stancik, & Fuller, 2003). The origins of protein film rheology in general have recently been discussed by Murray (2010). However, in this work we focus on the effect of added particles, which for the cellulose complexes this is to decrease significantly the temperature dependence of hi. Not surprisingly, this decrease is most marked for SC. As yet, the temperature dependence of gelatin þ complexes has not been studied, but overall it would appear that the addition of the complexes significantly changes the origins of the high film strength and that the increased strength is not particularly dependent upon hydrogen bonding interactions. Although the temperature range employed in the above experiments was rather narrow, it is still expected that the temperature variations will change the conformation of the adsorbed proteins to some extent, which may change the mechanism of interaction between the proteins and the complexes. Therefore, chemical methods of breaking hydrogen bonds were also tested, keeping the temperature constant. Fig. 11 shows the effect of including 6 mol dm3 urea in the solutions of 0.1 wt.% WPI or BL at the start of

-1

ΔG/ KJ mol 200 X

100 0 Fig. 9. Logarithm of the ratio of the interfacial shear viscosity (hi) at temperature T to that at 298 K versus (1/T) for: 0.1 wt.% whey protein isolate (;); 0.1 wt.% whey protein isolate þ 0.1 wt.% cellulose complexes (7); 0.1 wt.% b-lactoglobulin (C); 0.1 wt.% blactoglobulin þ 0.1 wt.% cellulose complexes (B); 0.1 wt.% sodium caseinate þ 0.1 wt.% cellulose complexes (>): using data after 24 h adsorption at the AeW interface at pH 6. Also shown are corresponding plots at pH 7 for: 103 wt.% sodium caseinate (-); 103 wt.% gelatin ().

0

2 -1 ηi / N s m

4

Fig. 10. Activation energy DG for flow obtained by fitting straight lines to the plots in Fig. 9 for the interfacial shear viscosity (hi) data below 30  C versus the absolute values of hi at 25  C after 24 h adsorption at the airewater interface (C). Also shown is the value obtained for gelatin for the data between 30 and 40  C ().

B.S. Murray et al. / Food Hydrocolloids 25 (2011) 627e638

ηi /

635

chemical reagents was observed for SC þ complexes, although it was hard to compare the effects of the treatments on SC alone because the hi values for SC were too low to be accurately measured with the experimental set-up used.

4

N s m-1 2

3.3. Starch particulates as stabilizing particles

0

0

1 t/ h

2

Fig. 11. Interfacial shear viscosity (hi) versus time (t) at the AeW interface at pH 6 for: 0.1 wt.% whey protein isolate þ 0.1 wt.% cellulose complexes (7); 0.1 wt.% whey protein isolate þ 0.1 wt.% cellulose complexes þ 6 mol dm3 urea (;); 0.1 wt.% blactoglobulin þ 0.1 wt.% cellulose complexes (B); 0.1 wt.% b-lactoglobulin þ 0.1 wt.% cellulose complexes þ 6 mol dm3 urea (C); 0.1 wt.% whey protein isolate þ 6 mol dm3 urea (,); 0.1 wt.% b-lactoglobulin þ 6 mol dm3 urea ().

the protein adsorption to the AeW interface at 25  C. The effect is to destroy any measurable hi for the whey proteins within less than 1 h. In contrast, when cellulose complexes are present along with urea, hi rises to significantly high values, albeit more slowly than without urea. This seems to confirm the conclusion from the temperature variation experiments that, hydrogen bonding between the protein and the complexes is not responsible for the high values of hi for the mixtures. Similar experiments were performed by adding 0.5 mol dm3 sodium thiocyanate to the WPI and BL systems. Urea disrupts both hydrogen bonding and hydrophobic bonding, through its effects on the hydrogen bonding structure of water, but at this concentration NaSCN is known to more specifically disrupt hydrophobic bonding (Damodaran & Kinsella, 1981) and Fig. 12 shows these effects. With no complexes present hi for WPI or BL is reduced to zero within 1 h by NaSCN, whereas in the presence of complexes hi continues to rise to significant values even in the presence of NaSCN. It should be pointed out that adding 0.5 mol dm3 NaSCN also significantly increases the ionic strength of the system, so that if attraction between opposite charges was important for the high values of hi for the proteins þ complexes, then the NaSCN might be expected to reduce hi via increased screening of electrostatic charges. Of course the addition of urea, sodium thiocyanate or other salts is expected to alter the conformation of the proteins, just as the temperature variation might. The additions may therefore also alter the interaction between the proteins and the complexes in complicated ways. Overall, however, one would expect at least one of the chemical or thermal treatments to affect the strength of the films significantly if hydrogen bonding, hydrophobic bonding or charge interactions between the particles and the whey proteins was important. Similar insensitivity of hi to the addition of the

The above experiments, using stable emulsion droplets or hydrophobic cellulose complexes as particles that can aid the stabilizing properties of proteins, have focused on the effects on bubbles and foams. Up to now we have not had much success using the cellulose complexes as O/W emulsifiers, probably because the particles are very much larger than the typical emulsion droplet sizes that one hopes to achieve for emulsions to remain stable to creaming, coalescence, etc. The cellulose particles are also mechanically strong and tend to lead to blockages of flow streams during attempts at emulsification. Nor have the emulsion droplet particles been investigated as emulsifiers themselves because this would require an oil phase that was immiscible with the droplet particles. There may be some non-polar liquids for which this is feasible, but these are unlikely to be food oils. For this reason the starch particulates have been investigated as emulsifiers, since the size and hydrophobicity of these particles was certainly easier to control than for the cellulose complexes. Some work is being been conducted on foam stabilization by more hydrophobic starch particulates than those described above, but this is not reported here. Fig. 13 shows a CLSM image of a 20 vol.% tetradecane emulsion stabilized solely by 3 wt.% of the hydrophobic starch particulates. Fluorescence from the added dyes has been colour coded so that that the starch particulates appear red and the oil phase appears yellow. Careful examination of such images reveals that all droplet surfaces are decorated by starch particulates and aggregates of particulates of widely varying size, although the coverage by clearly distinguishable particles is far from complete. It is well known (Binks, Clint, Mackenzie, Simcock, & Whitby, 2005; Binks & Horozov, 2006) that full coverage by surface active particles is not necessary to produce

2

ηi /

N s m-1 1

0 0

1

t/ h

2

3

Fig. 12. Interfacial shear viscosity (hi) versus time (t) at the AeW interface at pH 6 for: 0.1 wt.% whey protein isolate þ 0.5 mol dm3 NaSCN (;); 0.1 wt.% whey protein isolate þ 0.1 wt.% cellulose complexes þ 0.5 mol dm3 NaSCN (7); 0.1 wt.% blactoglobulin þ 0.5 mol dm3 NaSCN (C); 0.1 wt.% b-lactoglobulin þ 0.1 wt.% cellulose complexes þ 0.5 mol dm3 NaSCN (B).

Fig. 13. Typical confocal microscopy image of a 20 vol.% n-tetradecane emulsion stabilized by 3 wt.% surface active starch particulates. The fluorescence has been colour coded so that that the starch particulates appear red and the oil phase appears yellow. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

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B.S. Murray et al. / Food Hydrocolloids 25 (2011) 627e638

stable emulsions, as long as the adsorbed particle layer forms a rigid network. On the other hand, closer examination of the surfaces of larger droplets reveals a very thin, starch-rich region that appears to completely surround the droplets, although individual particles cannot be distinguished within it. This layer probably consists of a population of adsorbed starch particulates of sizes that are simply below the resolution of the microscope. It should be noted that extensive tests (Yusoff & Murray, 2011) have shown that there is no swelling or solubilisation of the starch particulates and that there are no other surface active molecular species present that could stabilize the emulsions. The system is therefore a true Pickering emulsion. Indeed, this is further substantiated by the high stability of the systems even though the mean droplet size is rather large, so that the systems creamed rapidly. Fig. 14 shows d32 over more than 6 months for emulsions stabilized by starch particulates that have been passed once through the homogenizer (homogenized starch) before making the O/W emulsion and also emulsions stabilized by non-homogenized starch particulates. The homogenized starch gave a smaller mean droplet size (d32 z 20 mm) than the non-homogenized starch (d32 z 30 mm), probably due to the greater surface coverage possible when the starch particle size is reduced by prior homogenization, but on re-dispersing both creamed emulsions d32 was shown to be essentially unchanged during 32 weeks storage. Finally, we report the interfacial shear viscosity of the starch particulate system at the n-tetradecaneewater interface, with and without added protein. Fig. 15 shows that hi for the (non-homogenized) 0.1 wt.% starch particle system alone is extremely low, as was also found for the cellulose complex particles at the airewater interface (Fig. 7). In separate experiments, not described here, we have found also that, on compressing an adsorbed film of the particles to as much as half the original film area, there is an increase in hi of less than 10%. Thus, as with the cellulose complexes as foam stabilizers, the starch particles are excellent emulsion stabilizers in their own right, but this stabilization is not dependent on high hi of the freely adsorbing film. Fig. 15 also shows what happens when 0.1 wt.% of the same starch particulates is combined with 0.1 wt.% SC. Again, as with the cellulose complexes, there is a remarkable increase in hi to values way above those found for either the starch or the SC on their own. The emulsifying properties of mixtures of proteins and surface active starch particles are currently being investigated, but it is tentatively suggested that significant enhancement in hi when surface active particles are combined with surface active proteins is a general effect that increases colloidal stability. Trying to account for the interfacial rheological effects of the particles when combined with proteins is seen to be difficult. Clues can possibly be obtained from studies by a number of workers on the interfacial rheology of systems of model surface active particles.

d32/ μm 30 20 10 0 0

10

20 t/ weeks

30

Fig. 14. Mean droplet diameter (d32) versus storage time (t) for 20 vol.% n-tetradecane emulsions stabilized by 3 wt.% surface active starch: homogenized particulates (>); non-homogenized particulates (-).

0.2

ηi/

N s m-1 0.1

0.0 0

10

t/ h

20

Fig. 15. Interfacial shear viscosity (hi) versus time (t) at the n-tetradecaneewater interface at pH 7 for: 0.1 wt.% surface active starch particulates þ 0.1 wt.% sodium caseinate (B); 0.1 wt.% surface active starch particulates only (C). The behaviour for 0.1 wt.% sodium caseinate on its own is indicated by the dashed line.

As indicated in the Introduction, many of these particles have no direct relevance to foods, but studies on model particles indicate some important underlying principles. Firstly, the aspect ratio of the particles is key. The packing of non-spherical particles at an interface is obviously very different from spheres and there is a strong tendency for the particles to become orientated at quite low fractions of occupancy of the interface. This is partly simply due to steric interactions, but also capillary interactions between particles. For example, Madivala, Fransaer, and Vermant (2009) have shown that the surface shear rheology of monolayers of ellipsoids is substantially greater than that of spheres even at low surface coverage. For such particles, shape-induced capillary (attractive) interactions compete with electrostatic (repulsive) interactions in determining the interfacial networks that form. Furthermore, the same types of particle can behave quite differently at AeW and OeW interfaces, due differences in these interactions. Madivala, Vandebril, Fransaer, and Vermant (2009) have shown that the stability of the corresponding emulsions is also strongly dependent on the particle aspect ratio and the resultant interfacial viscoelastic properties at a given surface coverage. Stable emulsions are only formed with particles above a certain aspect ratio. One key theoretical parameter is the maximum packing fraction for such systems. Basavaraj, Fuller, Fransaer, and Vermant (2006) have compressed monolayers of monodisperse prolate ellipsoidal latex particles in Langmuir trough experiments and shown that particles of a sufficiently large aspect ratio display a less abrupt increase surface pressure compared to spherical particles. Eventually, when the particles do jam, ellipsoidal particles of high aspect ratio can even flip into an upright position in the interface, unlike spherical particles. However, the restriction to monodisperse particles is somewhat unrealistic for practical systems. In theoretical studies, Delaney, Weaire, Hutzler, and Murphy (2005) and Groot and Stoyanov (2010) have illustrated how higher packing fractions can be achieved with polydisperse systems. Finally, although again based on monodisperse particles, Reynaert, Moldenaers, and Vermant (2006) have shown how interfacial aggregation and packing can be strongly affected by the presence of LMWS, due to their effects on the particle contact angle and therefore capillary interactions. Thus, the studies on model monodisperse particles at interfaces reveal that a rich range of behaviour is possible, even without the addition of proteins. As yet, few workers seem to have combined such model particles with even pure proteins. 4. Conclusions Surface active, food-grade particles have been prepared from cellulose and non-swelling starch that can act as effective stabilizers of foams and emulsions. When these particles are combined

B.S. Murray et al. / Food Hydrocolloids 25 (2011) 627e638

with proteins as surface active agents, there seems to be a synergistic effect in that the adsorbed film strength, as measured via the interfacial shear viscosity, is substantially increased and this increase seems to correlate with an increase in stability to coalescence and disproportionation. Protein-stabilized oil droplets can also produce similar effects at the airewater interface, but in this case the oil droplet ‘particles’ cannot be considered as actually adsorbing to the bare interface. In fact, with all the mixtures of particles and proteins used, it is not clear if both species are coadsorbing or if the particles adsorb at the surface of, or into, an adsorbed film of the protein. However, chemical and thermal treatment of the mixed films suggests that there are no specific weak interactions, such as hydrogen bonds or hydrophobic bonds, between the particles and the proteins that are responsible for the enhanced film strength. Possibly this points to enhanced incorporation of particles into the adsorbed layer in the presence of proteins that in turn leads to enhanced interfacial jamming of the particles at the interface. Further work on the structure and properties of such mixed protein þ particle films is required to substantiate this proposition, but it does appear that such mixtures could have an important role to play in the enhanced stabilization of food colloids in general. Acknowledgements The authors would like to thank EPSRC and Unilever Research and Development, Vlaardingen (URDV), The Netherlands for financial support of KD, the Ministry of Higher Education Malaysia and Universiti Teknologi MARA, Malaysia, for studentship funding of AY and in particular to Peter de Groot from URDV for useful discussions. The authors would also like to thank Erik van der Linden (Wageningen University) for useful discussions. References Allen, K. E., Dickinson, E., & Murray, B. S. (2008). Development of a model whipped cream: effects of emulsion droplet liquid/solid character and added hydrocolloid. Food Hydrocolloids, 22, 690e699. Allen, K. E., Murray, B. S., & Dickinson, E. (2008). Whipped cream-like systems based on acidified caseinate-stabilized oil-in-water emulsions. International Dairy Journal, 18, 1011e1021. Andresen, M., & Stenius, P. (2007). Water-in-oil emulsions stabilized by hydrophobized microfibrillated cellulose. Journal of Dispersion Science and Technology, 28, 837e844. Basavaraj, M. G., Fuller, G. G., Fransaer, J., & Vermant, J. (2006). Packing, flipping, and buckling transitions in compressed monolayers of ellipsoidal latex particles. Langmuir, 22, 6605e6612. Bhosale, R., & Singhal, R. (2006). Process optimization for the synthesis of octenyl succinyl derivative of waxy corn and amaranth starches. Carbohydrate Polymers, 66, 521e527. Binks, B. P., Clint, J. H., Mackenzie, G., Simcock, C., & Whitby, C. P. (2005). Naturally occurring spore particles at planar interfaces and in emulsions. Langmuir, 21, 8161e8167. Binks, B. P., Fletcher, P. D. I., & Holt, B. L. Phase inversion of nanoparticle-stabilised perfume oilewater emulsions: experiment and theory. Physical Chemistry Chemical Physics, in press, doi:10.1039/C0CP00558D. Binks, B. P., & Horozov, T. S. (2006). Colloidal particles at liquid interfaces. Cambridge: Cambridge University Press. Binks, B. P., Kirkland, M., & Rodrigues, J. A. (2008). Origin of stabilisation of aqueous foams in nanoparticleesurfactant mixtures. Soft Matter, 4, 2373e2382. Binks, B. P., & Lumsdon, S. O. (2000). Catastrophic phase inversion of water-in-oil emulsions stabilised by hydrophobic silica. Langmuir, 16, 2539e2547. Binks, B. P., & Rocher, A. (2009). Effects of temperature on water-in-oil emulsions stabilized solely by wax microparticles. Journal of Colloid and Interface Science, 335, 94e104. Borbas, R., Murray, B. S., & Kiss, E. (2003). Interfacial shear rheological behaviour of proteins in three-phase partitioning systems. Colloids and Surfaces A e Physicochemical and Engineering Aspects, 213, 93e103. Burgaud, I., Dickinson, E., & Nelson, P. V. (1990). An improved high-pressure homogenizer for making fine emulsions on a small scale. International Journal of Food Science and Technology, 25, 39e46. Campbell, A. L., Holt, B. L., Stoyanov, S. D., & Paunov, V. N. (2008). Scalable fabrication of anisotropic micro-rods from food-grade materials using an in shear

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flow dispersionesolvent attrition technique. Journal of Materials Chemistry, 18, 4074e4078. Campbell, A. L., Stoyanov, S. D., & Paunov, V. N. (2009a). Fabrication of functional anisotropic food-grade micro-rods with microparticle inclusions with potential application for enhanced stability of food foams. Soft Matter, 5, 1019e1023. Campbell, A. L., Stoyanov, S. D., & Paunov, V. N. (2009b). Novel multifunctional micro-ampoules for structuring and encapsulation. ChemPhysChem, 10, 2599e2602. Castle, J., Dickinson, E., Murray, B. S., & Stainsby, G. (1987). Mixed protein films at the oilewater interface. ACS Symposium Series, 343, 118e134. Cicuta, P., Stancik, E. J., & Fuller, G. G. (2003). Shearing or compressing a soft glass in 2D: timeeconcentration superposition. Physical Review Letters, 90, 236101. Cui, Z.-G., Yang, L.-L., Cui, Y.-Z., & Binks, B. P. (2010). Effects of surfactant structure on the phase inversion of emulsions stabilized by mixtures of silica nanoparticles and cationic surfactant. Langmuir, 26, 4717e4724. Damodaran, S., & Kinsella, J. E. (1981). The effects of neutral salts on the stability of macromolecules e a new approach using a proteineligand binding system. Journal of Biological Chemistry, 256, 3394e3398. Delaney, G., Weaire, D., Hutzler, S., & Murphy, S. (2005). Random packing of elliptical disks. Philosophical Magazine Letters, 85, 89e96. Dickinson, E. (2010). Food emulsions and foams: stabilization by particles. Current Opinion in Colloid and Interface Science, 15, 40e49. Dickinson, E., Ettelaie, R., Murray, B. S., & Du, Z. (2002). Kinetics of disproportionation of air bubbles beneath a planar airewater interface stabilised by food proteins. Journal of Colloid and Interface Science, 252, 202e213. Ettelaie, R., Dickinson, E., Du, Z., & Murray, B. S. (2003). Disproportionation of clustered protein-stabilized bubbles at planar airewater interfaces. Journal of Colloid and Interface Science, 263, 47e58. Groot, R. D., & Stoyanov, S. D. (2010). Equation of state of surface-adsorbing colloids. Soft Matter, 6, 1682e1692. Hotrum, N. E., van Vliet, T., Stuart, M. A. C., & van Aken, G. A. (2002). Monitoring entering and spreading of emulsion droplets at an expanding air/water interface: a novel technique. Journal of Colloid and Interface Science, 247, 125e131. Kostakis, T., Ettelaie, R., & Murray, B. S. (2007). Enhancement of stability of bubbles to disproportionation using hydrophilic silica particles mixed with surfactants or proteins. In E. Dickinson, & M. Leser (Eds.), Food colloids: Self-assembly and material science (pp. 357e368). Cambridge, UK: Royal Society of Chemistry. MacRitchie, F. (1986). Spread monolayers of proteins. Advances in Colloid and Interface Science, 25, 341e385. Madivala, B., Fransaer, J., & Vermant, J. (2009). Self-assembly and rheology of ellipsoidal particles at interfaces. Langmuir, 25, 2718e2728. Madivala, B., Vandebril, S., Fransaer, J., & Vermant, J. (2009). Exploiting particle shape in solid stabilized emulsions. Soft Matter, 5, 1717e1727. Marchessault, R. H., Morehead, F. F., & Walter, N. M. (1959). Liquid crystal systems from fibrillar polysaccharides. Nature, 184, 632e633. Maynard, A. D., Aitken, R. J., Butz, T., Colvin, V., Donaldson, K., Oberdorster, G., et al. (2006). Safe handling of nanotechnology. Nature, 444, 267e269. Moore, W. J., & Eyring, H. (1938). Theory of the viscosity of unimolecular films. Journal of Chemical Physics, 6, 391e394. Moschakis, T., Murray, B. S., & Biliaderis, C. G. (2010). Modifications in stability and structure of whey protein-coated o/w emulsions by interacting chitosan and gum arabic mixed dispersions. Food Hydrocolloids, 24, 8e17. Moschakis, T., Murray, B. S., & Dickinson, E. (2005). Microstructural evolution of viscoelastic emulsions stabilised by sodium caseinate and xanthan gum. Journal of Colloid and Interface Science, 284, 714e728. Murray, B. S. (1987). Pure and mixed protein films at the oilewater interface. PhD thesis, University of Leeds. Murray, B. S. (2002). Interfacial rheology of food emulsifiers and proteins. Current Opinion in Colloid & Interface Science, 7, 426e431. Murray, B. S. (2007). Stabilization of bubbles and foams. Current Opinion in Colloid and Interface Science, 12, 232e241. Murray, B. S. (2010). Rheological properties of protein films. Current Opinion in Colloid and Interface Science. doi:10.1016/j.cocis.2010.06.005. Murray, B. S., Cox, A., Dickinson, E., Nelson, P. V., & Wang, Y. (2007). Coalescence of expanding bubbles: effects of protein type and included oil droplets. In E. Dickinson, & M. Leser (Eds.), Food colloids: Self-assembly and material science (pp. 369e382). Cambridge, UK: Royal Society of Chemistry. Murray, B. S., Dickinson, E., Lau, C. K., & Schmidt, E. (2005). Coalescence stability of protein-stabilized bubbles when undergoing expansion at a simultaneously expanding planar airewater interface. Langmuir, 21, 4622e4630. Murray, B. S., Dickinson, E., & Wang, Y. (2009a). Bubble stability in the presence of oil emulsion droplets: influence of surface shear versus dilatational rheology. Food Hydrocolloids, 23, 1198e1208. Murray, B. S., Durga, K., de Groot, P. W. N., & Stoyanov, S. D. Preparation and characterization of the foam-stabilizing properties of celluloseeethyl cellulose complexes for use in foods, in preparation. Murray, B. S., & Ettelaie, R. (2004). Foam stability: proteins and nanoparticles. Current Opinion in Colloid and Interface Science, 9, 314e320. Murray, B. S., Færgemand, M., Trotereau, M., & Ventura, A. (1998). Comparison of the dynamic behaviour of protein films at the airewater and oilewater interfaces. In E. Dickinson, & J. M. Patino (Eds.), Food emulsions and foams: Interfaces, interactions and stability (pp. 223e235). Cambridge, UK: Royal Society of Chemistry. Murray, B. S., Xu, R., & Dickinson, E. (2009b). Brewster angle microscopy of adsorbed protein films at air-water and oil-water interfaces after compression, expansion and heat processing. Food Hydrocolloids, 23, 1190e1197.

638

B.S. Murray et al. / Food Hydrocolloids 25 (2011) 627e638

Paunov, V. N., Cayre, O. J., Noble, P. F., Stoyanov, S. D., Velikov, K. P., & Golding, M. (2007). Emulsions stabilised by food colloid particles: role of the particle adsorption and wettability at the liquid interface. Journal of Colloid and Interface Science, 312, 381e389. Pickering, S. U. (1907). Emulsions. Journal of the Chemical Society, 91, 2001e2021. Pradines, V., Kragel, J., Fainerman, V. B., & Miller, R. (2009). Interfacial properties of mixed beta-lactoglobulin-SDS layers at the water/air and water/oil interface. Journal of Physical Chemistry B, 113, 745e751. Ramsden, W. (1903). Separation of solids in the surface-layers of solutions and ‘suspensions’ (observations on surface-membranes, bubbles, emulsions, and mechanical coagulation). Preliminary account. Proceedings of the Royal Society, 72, 156e164. Reynaert, S., Moldenaers, P., & Vermant, J. (2006). Control over colloidal aggregation in monolayers of latex particles at the oilewater interface. Langmuir, 22, 4936e4945. Rousseau, D., Ghosh, S., & Park, H. (2009). Comparison of the dispersed phase coalescence methods in different tablespreads. Journal of Food Science, 74, E1eE7. Schmitt, C., Aberkane, L., & Sanchez, C. (2009). Proteinepolysaccharide complexes and coacervates. In G. O. Phillips, & P. A. Williams (Eds.), Handbook of hydrocolloids (2nd ed.). (pp. 420e476) Cambridge, UK: Woodhead. Schmitt, C., da Silva, T. P., Bovay, C., Rami-Shojaei, S., Frossard, P., Kolodziejczyk, E., et al. (2005). Effect of time on the interfacial and foaming properties of betalactoglobulin/acacia gum electrostatic complexes and coacervates at pH 4.2. Langmuir, 21, 7786e7795.

Turgeon, S. L., Schmitt, C., & Sanchez, C. (2007). Proteinepolysaccharide complexes and conjugates. Current Opinion in Colloid and Interface Science, 12, 166e178. Unterhaslberger, G., Schmitt, C., Shojaei-Rami, S., & Sanchez, C. (2007). Lactoglobulin aggregates from heating with charged cosolutes: formation, characterization and foaming. In E. Dickinson, & M. E. Leser (Eds.), Food colloids: Self-assembly and material science (pp. 177e194). Cambridge, UK: Royal Society of Chemistry. Wege, H. A., Kim, S., Paunov, V. N., Zhong, Q., & Velev, O. D. (2008). Long-term stabilization of foams and emulsions with in situ formed microparticles from hydrophobic cellulose. Langmuir, 24, 9245e9253. Whistler, R. L., & Paschall, E. F. (1967). Starch chemistry and technology. New York, London: Academic Press. Xu, R., Dickinson, E., & Murray, B. S. (2007). Morphological changes in adsorbed protein films at the airewater interface subjected to large area variations, as observed via by Brewster angle microscopy. Langmuir, 23, 5005e5013. Xu, R., Dickinson, E., & Murray, B. S. (2008). Morphological changes in adsorbed protein films at the oilewater interface subjected to compression, expansion and heat processing. Langmuir, 24, 1979e1988. Yusoff, A., & Murray, B. S. (2011). Modified starch granules as particle-stabilizers of oil-in-water emulsions. Food Hydrocolloids, 25, 42e55. Zhou, W. Z., Cao, J., Liu, W. C., & Stoyanov, S. (2009). How rigid rods self-assemble at curved surfaces. Angewandte Chemie-International Edition, 48, 378e381.