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Influence of the protein particle morphology and partition on the behaviour of particle stabilized water-in-water emulsions. Alberto Gonzalez-Jordan, Taco Nicolai, and Lazhar Benyahia Langmuir, Just Accepted Manuscript • Publication Date (Web): 23 Jun 2016 Downloaded from http://pubs.acs.org on June 23, 2016
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Influence of the protein particle morphology and partition on the behaviour of particle stabilized water-in-water emulsions.
Alberto Gonzalez-Jordan, Taco Nicolai, Lazhar Benyahia LUNAM Université du Maine, IMMM UMR-CNRS, Polymères, Colloïdes et Interfaces, 72085 le Mans Cedex 9, France. Abstract Protein fibrils, microgels and fractal aggregates were produced by heating solutions of -lactoglobulin (-lg) at different conditions. The effect of the protein particle morphology on the stability and the structure of water-in-water (W/W) emulsions was studied for mixtures of poly(ethylene oxide) (PEO) and dextran. The protein particles partition to the dextran phase at pH 7.0 where they have a net negative charge, but prefer the PEO phase at pH 3.0 where they have a net positive charge. The effect of partitioning on the stability and the structure of waterin-water (W/W) emulsions was studied by comparing emulsions at pH 3.0 with those at pH 7.0. The protein particle morphology and preference for one or the other phase are shown to have important consequences for the stability and the structure of the emulsions. Fibrils were found to be the most effective stabilizers at pH 7.0 whereas fractals were most effective at pH 3.0. The average droplet size obtained from confocal scanning laser microscopy was for most systems between 10 µm and 5 µm, but was notably smaller for emulsions with fractals at pH 3.0.
Introduction Water-in-water (W/W) emulsions are formed by mixing two incompatible water soluble macromolecules leading to a dispersed phase enriched with one macromolecule and a continuous phase enriched with the other1. The interfacial tension between the two phases is orders of magnitude smaller than for oil/water interfaces and is only expressed on length scales larger than the correlation length of the macromolecule solutions, i.e. several nanometers. In recent years stabilization of W/W emulsions using particles has attracted much attention 2-13. It has been shown that nano or micro particles can become irreversibly stuck at the interface and may inhibit fusion of the droplets. Particle stabilization has been known for a long time for oilin-water (O/W) emulsions and are known as Pickering emulsions14-17. When a particle enters the interface, the free energy is reduced by an amount that depends on their radius (R), the contact ACS Paragon 1Plus Environment
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angle with the interface () and the interfacial tension between the two phase separated aqueous macromolecule solutions (12):
G R 2 12 1 cos
2
1
The contact angle depends on the interaction of the particle with either one of the macromolecules that can be expressed in terms of effective interfacial tensions between the particles and each phase (p1, p2):
cos()=(p1-p2)/ 12
2
Recently, it was shown that not only spherical particles, but also disk-like18 and rod-like19 particles can stabilize W/W emulsions and can be explained by a similar mechanisms, but with different equations. A fourth type of particle that has not yet been tested are fractals that are self similar particles formed by random aggregation of elementary units20. Their shape is generally spherical, but their density () decreases with increasing radius (R) as R(3-df) with df the fractal dimension, which is most often situated between 1.6 and 2.3. The capacity of this type of particle to stabilize W/W emulsions has not yet been studied. Even though fractals are much less dense than microgels large polymers cannot penetrate the fractals to a significant extent so that it is expected that they will behave similarly to microgels. Of course, the contact angle of fractals with the interface will be less well defined as for smooth perfectly spherical particles. Interestingly three of these four different types of particles can be formed using the same globular protein. When globular proteins are heat-denatured they aggregate irreversibly and form microgels, fractal or fibrils depending on the conditions. The formation and the properties of the three types of aggregates have been studied in detail and has been reviewed 21-27 . When the net charge density () of the proteins is below a critical value they form homogeneous microgels whereas when is larger they form fractals with a df2 and a radius that increases with increasing protein concentration. The net charge of the proteins can be varied either by varying the pH or by adding multivalent ions. Finally, close to pH 2 the proteins hydrolyze and a fraction of the residual peptides assemble into rigid rod-like particles with lengths up to a few microns 28-31. The capacity of these three types of protein particles to stabilize O/W emulsions has already been investigated 32-33. However, a major difference with W/W emulsions is that native proteins can stabilize O/W emulsions in a manner similar to molecular surfactants, because they are amphiphilic and denature at the O/W interface. Native proteins cannot stabilize W/W ACS Paragon 2Plus Environment
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emulsions, because they are small compared to the correlation length of the macromolecule solutions. Generally protein aggregates solutions still contain a small fraction of native proteins or small denatured oligomers that will enter the O/W interface more quickly than the aggregates. In addition, the structure of the protein aggregates is modified by the interaction of the individual proteins with the O/W interface. Therefore it is not straightforward to interpret the effect of protein particle size and shape on the stabilization of O/W emulsions. Another important difference is that proteins are completely insoluble in the oil phase, whereas they are soluble in both water phases. However, in general the preference of proteins is not the same for two phases and they will partition in aqueous two phase systems 34-35. Our principal objective was to compare the capacity of protein fibrils, fractals and microgels to stabilize W/W emulsions against coalescence formed by mixtures of dextran and poly(ethylene) oxide (PEO). Protein particles with different morphologies were formed by thermal aggregation of -lactoglobulin (-lg), which is the mayor whey protein for which the isoionic point is at pH 5.0 36. In aqueous solution, stable suspensions of -lg microgels are formed in narrow pH ranges above and below the isoionic point (4.3-4.8 and 5.8-6.1) 37-39. Alternatively, microgels can be formed by heating -lg solutions at pH 7.0 by addition of CaCl2 37, 39-41 , which is the method used for this investigation. Electron microscopy images show that the microgels are approximately spherical and rather polydisperse. A fraction of proteins does not form microgels, but small curved strands with a length of approximately 50 nm and a diameter of approximately 10 nm 37, 40, 42-44. Microgels formed at pH 5.9 by heating whey protein isolate solutions flocculated in the pH range 4.0-5.8. Increasing the pH to 8 did not have a significant effect on the size, but decreasing the pH to pH 2 led to an increase of the hydrodynamic radius by about 20% 41. At pH>6.2, -lg forms small curved strands when heated in aqueous solution that at higher concentrations the strands randomly connect to form larger aggregates with a selfsimilar fractal structure 45. The size distribution of the fractals is broader than that of the microgels. In the pH range 1.5-2.5, -lg forms rigid-rod like aggregates 31, 46-48. The fibrils generally have a cross-section of a few nm and a distribution of contour lengths between 1 and several µm. The persistence length of the fibrils is about 1 µm 43, 52. Fractals formed at pH 7 and fibrils formed and pH 2 flocculate when the pH is set between pH 4 and pH 6. It has already been shown elsewhere8 that at pH 7.0 microgels of -lg spontaneously enter the PEO/dextran interface and that the layer of microgels on the surface of PEO droplets in the continuous dextran phase (P/D emulsions) inhibits coalescence of the droplets. Comparison of microgels with different average sizes showed that smaller microgels stabilized the P/D emulsions at lower protein concentrations. However, addition of native proteins had no noticeable effect on the rate at which the emulsions phase separated. The reason is that they were small compared to the correlation length of the of the polymers solutions so that the free energy does not significantly decrease when a native protein enter the interface. Addition, of ACS Paragon 3Plus Environment
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small -lg strands reduced weakly the rate of macroscopic phase separation, indicating that the strands entered the interface, but could not arrest coalescence. It follows that the stabilization of W/W emulsions by large protein aggregates is not significantly influenced by residual native proteins or small strands. This is an important difference with O/W emulsions for which residual native proteins, peptides or strands accumulate more rapidly at the interface and effectively compete with larger aggregates. Interestingly, it was found that dextran droplets covered by a layer of microgels in the continuous PEO phase (D/P emulsions) were not stable contrary to PEO droplets in P/D emulsions. A continuous bottom layer of dextran containing microgels was formed rapidly. Here we will compare these results with those obtained with fibrils and fractals. A second objective was to investigate the effect of the net charge of the proteins and their preference for each phase by comparing emulsions at pH 7.0 with emulsions at pH 3.0. We will show that the proteins prefer the dextran phase at pH 7.0 and the PEO phase at pH 3.0.
Materials and methods Formation and characterization of protein particles The β–lg (Biopure, lot JE 001-8-415) used in this study was purchased from Davisco Foods International, Inc. (Le Sueur, MN, USA) and consisted of approximately equal quantities of variants A and B. The powder was dissolved in pure water (Millipore) containing 200 ppm NaN3 to protect against bacterial growth. The solutions were filtered through 0.2 µm pore size filters (Anatope) and the pH was set by the addition of aliquots of 0.1 M HCl or 0.1 M NaOH under vigorous stirring. Fractals and microgels were prepared by heating the protein solutions at pH 7.0 overnight at 80°C. After this heat treatment the fraction of residual native proteins was negligible. More than 90% of the proteins form fractals after heating in pure water at C=95 g/L 45 and more than 80% formed microgels at C= 40 g/L in the presence of 4.4 mM CaCl253. In solutions of fractals the remaining proteins consisted of denatured monomers and small oligomers 45, whereas solutions of microgels also contained small strands. Fibrils were prepared by heating protein solutions at pH 2.0 and C= 20 g/L for 5 h at 90°C under stirring as described in ref. 52. It was found that conversion of -lg into fibrils can be accelerated by stirring 49-51. The fibrils had a cross section of 5 nm and lengths between 1 and 20 µm. By measuring the sedimentation at pH 4.6 it was found that 75% of the proteins where aggregated. However, it is possible that a smaller fraction had actually formed fibrils 47. The remaining proteins consisted of unassociated peptides and residual native proteins. The protein concentration was determined by measuring the adsorption of UV-light with wavelength 280 nm (Varian Cary-50 Bio, Les Ulis, France) using and extinction coefficients of 0.96 L g−1 cm−1. The z-average hydrodynamic radius of the fractals and the microgels was ACS Paragon 4Plus Environment
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determined using dynamic light scattering as described in ref.53 and was found to be 150 nm. As was mentioned in the introduction neither the microgels nor the fractals are monodisperse in size and in particular the fractals have a broad size distribution. Notice that the z-average hydrodynamic radius gives strong weight to the larger aggregates. Transmission electron microscopy images of the fibrils showed they were several µm long and had a diameter of about 5nm52. All three types of particles aggregated between pH 6 and pH 4. However, if the pH of the fibril solution was increased from pH 2.0 to pH 7.0 by rapidly adding the required amount of NaOH while stirring aggregation of the fibrils could be avoided. Elsewhere it was shown that long fibrils persist after increasing the pH to 7.0 54 55, but it is likely that the average length of the fibrils was reduced. In the same manner the pH of the microgels and the fractals could be decreased from pH 7.0 to pH 3.0 without aggregation of the particles. Fractals and microgels are sufficiently robust to remain intact during this process. It was reported elsewhere that the size of the microgels varies little between pH 7.0 and pH 3.0 54. We found using dynamic light scattering that the hydrodynamic radius of the fractal was also the same at the two pH values. Preparation of the emulsions Dextran and PEO were purchased from Sigma-Aldrich and dissolved under stirring in Millipore water. The weight average molar mass was Mw = 1.6 × 105 g/mol for the dextran and Mw = 2 × 105 g/mol for the PEO. The PEO powder contained a small amount of silica particles that was removed by centrifugation of the PEO solutions at 5x104 g during 4h. Emulsions were prepared by mixing aqueous solutions of PEO, dextran and protein particles at pH 7.0 in the required amounts using a minishaker. Neither the order of the addition of the components nor the stirring speed or duration had a significant influence on the structure or behaviour of the emulsions. The phase diagram of mixtures of the PEO and a dextran samples used here is shown in fig. 1. We found that the binodal did not depend on the pH.
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5 4 3 2 1 0 0
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Cdex (wt%)
Fig. 1 Phase diagram for mixtures of the PEO and dextran samples used for this study. The open and filled circles represent homogeneous and phase separated mixtures, respectively. The solid line represents the binodal and the dashed-dotted lines represent tie-lines. The red square and the blue triangle represent the mixtures that were investigated here. Confocal microscopy The proteins particles were visualized with a confocal laser scanning microscope (CLSM) by addition of 5 ppm rhodamine to the solutions which binds spontaneously to the proteins. CLSM observations were made with a Leica TCS-SP2 (Leica Microsystems Heidelberg, Germany). Images of 512 × 512 pixels were produced at different zooms with two different water immersion objectives: HC× PL APO 63× NA = 1.2 and 20× NA = 0.7. The solutions were inserted between a concave slide and a cover slip and hermetically sealed. The incident light was emitted by a laser beam at 543 nm and/or at 488 nm. The fluorescence intensity was recorded between 560 and 700 nm. It was verified that the use of labeled dextran and proteins had no influence on the emulsions.
Results and discussion Emulsions at pH 7. P/D emulsions were prepared by mixing 1.9 % (w/w) PEO, 12% dextran (red square in fig. 1) and different quantities of protein particles between C=0.05 % and 0.75 % leading to formation of droplets of the PEO phase with a volume fraction of 25 % in the continuous dextran phase. At these conditions the polymers practically fully phase separated with 8.2 % ACS Paragon 6Plus Environment
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PEO in the droplets and 15.8 % dextran in the continuous phase. The interfacial tension between the two phases at these conditions was found to be 75µN/m 2. Fig. 2 shows photographs of the solutions after 1 day, 3 days and 1 week. In the presence of microgels the emulsions formed a clear top layer of the PEO phase, which implies that coalescence of the droplets had occurred. The rate at which the PEO droplets coalesced into a macroscopic PEO phase decreased with increasing protein concentration and was insignificant at least for one week for C0.5%. However, even if the proteins particles can inhibit coalescence completely they cannot stop the PEO droplets from creaming and forming a dense droplets layer at the top. Elsewhere we showed that the rate of creaming could be qualitiatively explained on the basis of the average size of the droplets, the difference in density between the two phases and the viscosity of the continuous phase8. However, quantitative agreement was not obtained, because the effects of droplet size dispersity and interaction between creaming droplets was not considered. The same calculations can be applied to creaming of droplets stabilized by fractals or fibrils, because the effect of the thin interfacial layer on the density of the droplets is negligible. With increasing protein concentration the turbidity of the homogeneous bottom dextran layer increased, because excess microgels preferentially migrated to the dextran phase at pH 7.0, see below. Emulsions containing fractals formed a clear top layer of the PEO phase much faster than those with microgels and a layer was visible after 1 week standing even at the highest protein concentration investigated. Remarkably, emulsions that contained 0.1 % fractals were much more stable than either at C=0.05 % or C=0.2 %. Here we consider emulsions stable if the droplets do not coalesce even if they cream or sediment. This effect was reproducible and emulsions at C=0.075 % and C=0.15 %, were also significantly more stable than at higher or lower protein concentrations. We have currently no explanation for this intriguing behavior. Fibrils were found to be the most effective stabilizers of the emulsions at pH 7. Even at C= 0.05 % formation of a clear PEO top phase was very slow. Interestingly, creaming of the PEO droplets was also much slower in the presence of fibrils and was almost absent after one week at C=0.3 %. It was shown elsewhere 8 that the same behaviour could be obtained with smaller microgels than the ones used here, but only at C> 0.5 %. The effective stabilizing effect of the fibrils is all the more remarkable if we consider that it contains a significant fraction of proteins that did not form fibrils. We note that since the fibrils have a persistence length of about 1 µm so that there is no enthalpic penalty to cover the surface of droplets with radii that are larger than 1µm, which is the case here, see below.
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Microgels
Fractals
Fibrils 1 day
3 days
1 week
Fig. 2. Photographs at different times after mixing of PEO in dextran emulsions containing different amounts of microgels or fractals: 0.05, 0.1, 0.2, 0.3, 0.5, 0.75 % (from left to right), or fibrils: 0.05, 0.1, 0.3 % (from left to right).
CLSM images of the P/D emulsions just after mixing are shown in fig. 3. Fluorescence of rhodamine B labeled proteins is indicated by the red coloring. In mixtures with the microgels a layer of proteins covering the PEO droplets can be clearly distinguished. Excess protein particles go preferentially to the dextran phase at pH 7.0, which explains the stronger fluorescence intensity of the dextran phase at high protein concentrations. However, residual unbound rhodamine B partitions preferentially to the PEO phase, which explains why this phase is more fluorescent at low protein concentrations. A few larger aggregates of can be seen in the solutions containing microgels, which is probably caused by depletion interactions between the polymers and the microgels. In the presence of fractals or fibrils the protein layer covering the PEO droplets is not clearly visible even though the increased stability of the emulsions shows that it must be present. Only at the lower protein concentrations can a faint interfacial layer of fractals be seen in the images. The reason that the layer cannot be clearly seen is that the density of the proteins at the interface is much lower for the fractals and the fibrils. Fractals are spherical, but much less dense than the microgels, whereas the fibrils are most likely oriented parallel to the interface leading to a very thin layer. The number average radius of the droplets (R) decreased weakly with increasing protein concentration from 107 µm at 0.05 % to 64 µm at 0.75 % for the microgels, from 85 µm at 0.05 % to 53 µm at 0.75 % for the fractals and from 74 µm at ACS Paragon 8Plus Environment
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0.05 % to 64 µm at 0.3 % for the fibrils. The concentration of excess protein aggregates in the continuous dextran phase is very small and therefore is not expected to have a significant influence on the viscosity of the dextran phase. Indeed, we did not observe a significant difference in the flow of tilted emulsions. Since the morphology of the protein particles does not have a strong effect on the initial droplet size, one would therefore expect that the rate of creaming was similar. The significantly slower creaming for emulsions stabilized by fibrils is most likely caused by reduced coalescence during creaming of the droplets stabilized by fibrils. Nevertheless, it is possible that the highly asymmetric fibrils did influence the effective viscosity felt by the droplets that exert very low stress and that cannot be observed when the samples are tilted. This would explain the strong effect of the fibril concentration on the creaming rate.
Microgels
Fractals
Fibrils 0.05%
0.1%
0.3%
0.75%
Fig 3. CLSM images (160x160 µm) of PEO in dextran emulsions in the presence of different concentrations of microgels, fractal or fibrils. Proteins are labeled with rhodamine B, but residual unbound rhodamine B is preferentially situated in the PEO phase.
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Emulsions of droplets of the dextran phase with a volume fraction of 25% in the continuous PEO phase were prepared by mixing 6.3 % PEO and 4.0 % dextran (blue triangle in fig. 1). The composition of the mixture was situated on the same tie-line tie line as the PEO in dextran emulsion so that the interfacial tension was the sa same. me. It was already shown elsewhere8 that D/P emulsions were not stabilized by microgels. The same behavior was found here for emulsions with fractals or fibrils. fibrils Fig. 4 shows that even though a layer of microgels can be observed at the interface, the droplets coalesce rapidly into llarge arge dextran domains.
50 µm
100 µm
Figure 4. CSLM images of droplets roplets of the dextran phase taken at the bottom of a dextran in PEO emulsion in the presence of 0.3 % microgels at pH 7.0 immediately after mixing (left) and after 3 h (right). Notice that the scale of the images is different.
Emulsions at pH 3. Similar experiments were done at pH 3.0. Fig. 5 shows photographs of the PEO in dextran (P/D) emulsions at different times time after preparation. The behavior of the emulsions at pH 3.0 was strikingly different from that at pH 7.0. At pH 3.0, fibrils did not stabilize PEO droplets at all, while with fractal aggregates or microgels the emulsions were stable for at least 1 week. Creaming of PEO droplets was very fast for emulsions containing microgels, whereas it was very ry slow when fractals were present. The observed fast creaming of PEO droplets covered with microgels corresponds to the t fast sedimentation of microgel covered dextran droplets at pH 7.0. Both are caused by agglomeration of the droplets into larger clusters cluster that cream or sediment rapidly. Notice that at pH 3.0 the continuous dextran bottom layer is clear for the mixtures with microgels and that the continuous PEO top layer is turbid. turbid This means that excess microgels partitioned preferentially to the dextran phase at pH 3.0 and not to the dextran layer as was the case at pH 7.0.
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Microgels P/D
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Fig. 5. Photographs at different times after mixing of PEO in dextran (P/D) emulsions and dextran in PEO (D/P) emulsions with different amounts of microgels, fractals or fibrils: 0.05, 0.1, and 0.3 % (from left to right). Another major difference with emulsions at pH 7.0 is that at pH 3.0 stable emulsions could be formed of dextran droplets in the continuous PEO phase (D/P), whereas at pH 7.0 D/P emulsions destabilized rapidly. Dextran droplets could be stabilized at pH 3.0 with all 3 types of protein particles and sedimented slowly. However, the rate at which the droplets sedimented depended strongly on the type of particle. Dextran droplets covered with fibrils sedimented more slowly than those covered with microgels and droplets covered with fractals sedimented much more slowly than those covered with either micorgels or fibrils. The turbidity of the PEO top phase in mixtures with microgels shows that it contained most of the excess microgels. CLSM images of the emulsions just after mixing are shown in fig.6. The PEO droplet radius distribution of P/D emulsions was similar to that at pH 7.0 in the presence of microgels (R=75 µm) or fibrils (R=75 µm) and did not vary much with the protein concentration at least ACS Paragon11 Plus Environment
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between 0.05 and 0.3%. However, in mixtures with fractals the droplets were significantly smaller and the size decreased with increasing protein concentration from R=53 µm at 0.05 % to R=2.51.5 µm at 0.3 %. This explains why creaming of PEO droplets was much slower for mixtures with fractals. The size of the dextran droplets in D/P emulsions with microgels (R=4.52.5 µm) or fibrils (R=3.53 µm) did not vary much with the protein concentration and was smaller than that of the PEO droplets shown in fig. 6. The smallest droplets were again formed with the fractals and the average radius decreased with increasing protein concentration from 2.51.5 µm at 0.05 % to 10.5 µm at 0.3 %.
Microgels P/D
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Fig 6. CLSM images (160x160 µm) of PEO in dextran and dextran in PEO emulsions in the presence of different concentrations microgels, fractals fractal or fibrils. The insert in one of the images represents a zoom (20x20 µm).
The effect of ageing was studied for emulsions containing contain g 0.3 % protein particles. Figure 7 shows images of aged P/D and D/P emulsions in the presence of microgels taken near the top and the bottom of the samples, respectively. PEO droplets coalesced into larger droplet and therefore creamed rapidly, but they did not fuse in to a homogeneous PEO layer. Dextran droplets remained smaller and sedimented more slowly. In the presence of fractals the droplet size did not change significantly with ageing, whi which ch explains the slow creaming and sedimentation. In the presence of the fibrils PEO droplets coalesced and creamed rapidly into a homogeneous PEO top phase. The size of the dextran d droplets did not increase significantly, which explains why they sedimented slowly.
Fig. 7. CSLM images (160 µm x160 µm) of droplets of PEO phase taken at the top of a PEO in dextran emulsion (left) and of droplets of the dextran phase taken at the bottom of a dextran in PEO emulsion. The emulsions were pH 3.0 and contained 0.3 % microgels. microgels. The images were taken 2 days after preparation.
Partitioning of proteins in the two phases It was already noted that excess microgels partitioned to the dextran phase at pH 7.0 and to the PEO phase at pH 3.0. Partitioning of proteins in mixtures of dextran and PEO has been studied in detail 34-35. It depends on the type of protein, but also on the polymer concentration, molar mass and the pH. W Wee have determined the partition coefficient K defined as the ratio of the protein n concentration in the PEO phase to that in the dextran phase, as described in the materials and methods section. Fig. 6 shows that K has a minimum value at pH 5.0, i.e. close to the isoionic point of the proteins. A minimum value of K is expected at th the isoionic point,, because the electric potential opposes partitioning of charge particles. This effect ACS Paragon13 Plus Environment
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is expected to be reduced by addition of salt. Indeed, we observed a decrease of K to K=0.14 when 0.1 M NaCl was added to the solutions at pH 7.0. However, addition of salt not only influences the partitioning, but also introduces attractive interaction between the aggregates which is known as cold gelation 56. The effect of cold gelation on W/W emulsions stabilized by protein particles will be discussed elsewhere. The effect of the ph on the net charge density of the proteins cannot explain why K became larger than unity for pH
