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Effect of protein aggregates on foaming properties of β-lactoglobulin
Colloids and Surfaces A: Physicochem. Eng. Aspects 330 (2008) 96–102
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Effect of protein aggregates on foaming properties of -lactoglobulin Bénédicte Rullier, Bruno Novales ∗ , Monique A.V. Axelos Institut National de la Recherche Agronomique, UR1268 Biopolymères, Interactions, Assemblages, INRA, Equipe Interfaces Systèmes Dispersés, Rue de la Géraudière, 44316 Nantes Cedex 03, France
a r t i c l e
i n f o
Article history: Received 3 June 2008 Received in revised form 16 July 2008 Accepted 19 July 2008 Available online 30 July 2008 Keywords: Foam Protein aggregates Interfacial properties
a b s t r a c t Our paper aims at determining the respective part of protein aggregates and non-aggregated proteins in the foam formation and stability of -lactoglobulin. We report results on fractal aggregates formed at neutral pH and strong ionic strength (aggregates size from 30 to 190 nm). Pure aggregates and mixtures of non-aggregated/aggregated proteins at varying ratios were used. The capacity of aggregates to form and stabilize foams has been studied in relation with their ability to absorb at air/water interfaces. Our results show that protein aggregates are not able by themselves to improve the foaming properties but participate to a better foam stabilization in the presence of non-aggregated proteins. Non-aggregated proteins appear to be necessary to produce stable foams. We have shown that the amount and the size of aggregates had an influence on the drainage rate. © 2008 Elsevier B.V. All rights reserved.
1. Introduction Aqueous foams are dispersions of gas bubbles in a liquid stabilized by amphiphilic molecules [1]. In the food industry, foams are stabilized mainly by proteins. A large number of papers have been devoted to the characterization of the foaming behavior of proteins under a variety of conditions [2,3]. Surfactants rapidly decrease the surface tension to low values, while proteins slowly but strongly adsorb to the air/water interface. Even if the decrease of surface tension is less important for proteins, it has been demonstrated to be correlated to their foaming ability [4–6]. In particular, it has been shown that flexible proteins with high surface activity adsorb more rapidly to the interface than compact globular proteins [7]. When adsorbed at the interface, proteins create high viscoelastic layers with high surface shear viscosities. Martin et al. [7] have been demonstrated that under certain conditions, the adsorbed protein film can form a resistant and cohesive interfacial network which enhances foam stability. Most of the studies cited have been carried out on nonaggregated proteins. However, during many food processings, proteins undergo thermomechanical treatments that often lead to their aggregation in self-assembled structures [8–10]. As protein aggregates are complex structures whose size may be as large as a few hundred nanometers, it can be expected that their behavior is different from non-aggregated proteins. Because of their size, the ability of aggregates to diffuse to the interface is reduced and this
∗ Corresponding author. Tel.: +33 240675198; fax: +33 240675084. E-mail address: [email protected] (B. Novales). 0927-7757/$ – see front matter © 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.colsurfa.2008.07.040
should lead to lower foaming properties. Moreover, protein–protein interactions within aggregates can be of a different nature (electrostatic, hydrophobic, etc.), leading to different kinds of cohesiveness. This could affect the mobility of proteins and their ability to spread at the interface and influence their capacity to stabilize air/water interfaces needed to produce a foam. However, it has been shown that the presence of some aggregates increases the foaming stability [9,11,12]. The mechanism by which aggregates contribute to foam stabilization is not completely elucidated. Different hypotheses have been proposed: either the protein aggregates can adsorb at the interface or they cannot. If they do, aggregates increase the viscoelasticity of the interface thus providing a better foam stabilization [12]. On the other hand, if aggregates do not adsorb to the interface, they remain in the aqueous phase. In this case, aggregates can become confined into foam films and can undergo a percolation process leading to the formation of a gel-like network. They can also prevent foam destabilization by reducing drainage acting as cork in the Plateau borders [13]. In industrial processes, only a given fraction of the proteins is aggregated and the part of non-aggregated proteins may considerably influence the foaming properties. Therefore, it is often difficult to determine the intrinsic role of aggregates in these systems. The aim of our paper is to determine the respective part of protein aggregates and of non-aggregated proteins played in the foam formation and foam stability of protein solutions. For the present work -lactoglobulin has been chosen because its aggregation properties in function of experimental parameters such as temperature, pH, ionic strength and protein concentration have been studied in considerable detail [14–16]. The size and the shape of -lactoglobulin aggregates can be easily varied by the choice of the experimen-
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tal conditions, leading to fibrillar or fractal aggregates with sizes ranging from a few nanometers to hundreds of nanometers. In this paper, we report results on fractal aggregates exclusively formed by heating at 80 ◦ C during 24 h at neutral pH and 0.1 M of ionic strength. The protein concentration was varied in order to modulate the size of the aggregates. The capacity of the aggregates to form and stabilize foams has been studied in relation with their ability to absorb at air/water interfaces.
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The amount of non-aggregated protein after the UF–C step was determined by gel filtration chromatography (GFC) under the following conditions: column TSK PW 6000 equilibrated with an elution solution composed of Trizma NH2 C(CH2 OH)3 0.05 M; NaCl 0.1 M at pH 7. The eluted protein was detected using UV adsorption at 280 nm and the amount of residual non-aggregated protein was calculated as percent area relative to the area of unheated protein. 2.3. Surface tension measurements
2. Materials and methods 2.1. ˇ-Lactoglobulin -Lactoglobulin was purified from whey protein isolate (Prolacta 90 batch no. 273 supplied by Lactalis Industrie, Laval, France) according to the method of Maillart and Ribadeau-Dumas [17]. After lyophilisation, -lactoglobulin was analyzed by mass spectrometry. Each sample of -lactoglobulin was constituted of 55% of -IgA, 35% of -IgB and 10% of -IgA lactosylated. The average mass was determined at 18,385 kDa. -Lactoglobulin powder was dissolved in milliQ water at pH 7 containing 3 mM sodium azide to prevent from bacterial contamination. -Lactoglobulin solution was dialyzed and filtered with a 0.22 m membrane. Protein solutions were then dissolved in a 0.1 M sodium chloride (NaCl) solution at pH 7. Protein concentrations were determined through the measurement of UV adsorption at 278 nm, taken an extinction coefficient of 0.96 L g−1 cm [16]. 2.2. Formation and characterization of protein aggregates -Lactoglobulin aggregates were formed upon heat-induced denaturation of proteins at different concentrations at a temperature of 80 ◦ C during 24 h. The samples were cooled in a bath at room temperature. In this paper, aggregates were formed at pH 7 and 0.1 M NaCl ionic strength at four concentrations: 1, 2, 8 and 10 g/L. Aggregate sizes were determined by static and dynamic light scattering. Static measurements were made using an ALV-5000 multi-bit multi-tau correlator and a Spectra Physics solid-state laser operating with polarized light with wavelength = 532 nm. The range of scattering wave vectors covered was from 3.0 × 10−3 to 3.5 × 10−2 nm−1 (q = 4ns sin(/2)/, with the angle of observation and ns the refractive index of the solution). Dynamic light scattering was performed using a Nanosizer ZS (Malvern Instruments, UK) in order to determine the hydrodynamic radius of the aggregates. The instrument was used in the backscattering configuration where detection is done at a scattering angle of 173◦ . Dilute solutions were measured in a 1-cm path-length spectroscopic plastic cell at 20 ◦ C. The hydrodynamic radius was measured in triplicate. Each measurement corresponded to three autocorrelation functions recorded during 90s. As the size of our aggregates were close to the range of reliability of the Malvern Nanosizer (given by the manufacturer), data obtained from this apparatus were compared to results obtained by static light scattering for some samples in order to check the reliability of our measurements. In order to obtain solutions containing exclusively protein aggregates, the non-aggregated proteins remaining in the solution were separated out by an ultrafiltration–centrifugation (UF–C) using an Amicon membrane Ultra 30 kDa. This step was performed during 30 min at 4000 rpm and 4 ◦ C for the larger aggregates, and during 4 h for the smaller ones. The retentate was redispersed in a 0.1 M NaCl solution and the concentration was verified by the measurement of UV absorbance at 278 nm.
An automated drop tensiometer (IT-Concept, Longessaigne, France) was used to measure the interfacial tension between liquid and air, as described in detail by Benjamins et al. [18]. All experiments were performed in the rising bubble configuration. A 6 L air bubbles was formed upward at the tip of a U-shaped stainless steel needle immersed in a cuvette filled with the protein solution, under gentle stirring, both of which are temperature controlled at 20 ◦ C. The dynamic tension was followed in time (120 s in our case) using axisymmetric drop-shape analysis. Image acquisition and regression of the interfacial tension were performed with Windrop software by fitting the Laplace equation to the drop shape. 2.4. Foam experiments Foaming experiments were conducted on a “Foamscan” apparatus developed by IT Concept (Longessaigne, France). With this instrument, the foam formation, the stability and the drainage of liquid from the foam can be determined by conductimetric and optical measurements. After calibration of the conductimetric electrodes, foams were produced from 12 mL of the solution at protein concentration 1 g/L by injection of gaseous N2 through a porous glass filter. The flow rate was fixed at 35 mL/min. All foams were allowed to reach a final volume of 45 mL after which gas-flow was stopped and the evolution of the foam was analyzed. The foamability corresponds to the time needed to reach 45 mL of foam volume. After foam formation, in a timespan of 3600 s, several parameters are automatically recorded by the “Foamscan” analyzing software. The volume of the foam is determined with a CCD camera (Sony Exwave HAD). A pair of electrodes at the bottom of the column is used to measure the quantity of liquid that is not in the foam, while the volume of the liquid in the foam is measured by conductimetry using three pairs of electrodes located along the glass column. The liquid drainage of the foam is followed via conductivity measurements at different heights in the foam column. In order to get information about the composition of the protein solutions and foams, the drained liquid was quantified using the GFC method previously described in detail. These measurements were made at two different times: firstly just after the foam formation when the desired foam volume is reached and secondly 1200 s after the end of bubbling. For long-time drainage experiments (3600 s), in order to check that the drainage was not affected by other destabilization mechanisms (such as coarsening), some foams were produced by injecting N2 through a perfluorohexane (C6 F14 ) solution. We observed that at the electrode 1 (where the liquid fraction is measured), the drainage curves had similar profiles with or without C6 F14 . 3. Results 3.1. Characterization of protein aggregates From static light scattering measurements, the dependence of the scattered intensity Ir (q) on the scattering wave vector (q) by the aggregates at different protein concentrations can be obtained.
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Fig. 2. Dynamic surface tension measurements of non-aggregated proteins and protein aggregates at C = 0.05 g/L: (䊉) non-aggregated protein; aggregates () 35 nm; () 71 nm; (×) 117 nm; ()197 nm.
the rest of this paper, when referring to protein aggregate samples their hydrodynamic radius measured after UF–C is meant. Fig. 1. Master curves of normalized scattering wave vector (q) as function of the normalized intensity of scattered light (Ir ) for dilute solutions of -lactoglobulin aggregates.
From the initial q-dependence the z-average radius of gyration (Rgz ) of the aggregates can be extracted. After dividing Ir (q) by the weight average molar mass Mw and plotting the result as a function of qRgz , a master curve is obtained (Fig. 1) which means that aggregates are self-similar. At high qRgz -values, we observe a power law dependence of Ir (q) ∞ q−df . The exponent df represents the socalled fractal dimension of the aggregates [16]. In our case, df is close to 2 which corresponds to typical fractal aggregates. From the weight average molar mass Mw determined from the curves, it is possible to determine the average number of proteins within aggregates. The measured number ranges from a few hundred proteins for the small aggregates to around 5000 proteins for the larger ones. From dynamic light scattering the size distribution of nonaggregated proteins as well as of protein aggregates was obtained. Non-aggregated -lactoglobulin has a hydrodynamic radius of 3 nm and the weight molar mass Mw was determine at 33 kg/mol, which is coherent with data in the literature [15]. The average hydrodynamic radius (Rh ) of protein aggregates are reported in Table 1. The aggregate size increases with protein concentration with Rh ranging from 35 nm at 1 g/L to 197 nm at 10 g/L. By contrast, the amount of non-aggregated protein, determined by GFC, increases when the protein concentration decreases. When the heat treatment is stopped after 24 h, the equilibrium has not yet been reached and as the aggregation process depends on time and protein concentration, this explains the observed variations of aggregate size and aggregates/non-aggregated protein ratios. In order to obtain exclusively protein aggregates, nonaggregated proteins were separated from the aggregates by UF–C. After the UF–C step, the aggregate size was checked by dynamic light scattering, and the amount of non-aggregated proteins was determined by GFC to be close to 0.5% for large aggregates (117 and 197 nm) and close to 1% for small aggregates (35 and 71 nm). For
3.2. Surface tension As foam formation occurs in a few seconds, it is more pertinent to study the adsorption kinetics at short times when attempting to explain the ability of proteins to adsorb at air/water interfaces. The surface tension of both non-aggregated proteins and protein aggregates measured during 120 s is shown in Fig. 2. For non-aggregated -lactoglobulin, we observed the fastest decrease of the surface tension, with a value lower than 60 mN/m after 120 s. For protein aggregates, the adsorption kinetic is slower. The adsorption rate of the aggregates decreases with their size. Three groups of adsorption rate behavior can be distinguish: (i) for the largest protein aggregates no decrease of the surface tension was observed; (ii) intermediate size aggregates with Rh 71 and 117 nm; and (iii) small protein aggregates (Rh 35 nm) which, during the first 10 s, present a decrease of surface tension that is similar to that of non-aggregated proteins. 3.3. Foaming properties The time evolution of the foam volume for non-aggregated proteins and for protein aggregates is shown in Fig. 3. A foam made from non-aggregated proteins is formed in 82 s. Such a foam is relatively
Fig. 3. Time evolution of foam volume for non-aggregated proteins and proteins aggregates solutions (1 g/L).
Table 1 -Lactoglobulin aggregates hydrodynamic radius and fractions of non-aggregated proteins measured before and after UF–C step as function of protein concentration Protein concentration (g/L)
Before UF–C Rh (nm)
1 2 8 10
34 66 111 196
± ± ± ±
6 1 1 5
After UF–C % of non-aggregated proteins 20.4 10.4 10.6 3.7
± ± ± ±
10.0 2.0 2.0 0.6
Rh (nm) 35 71 117 197
± ± ± ±
1 1 1 10
% of non-aggregated proteins 1.0 0.80 0.80 0.50
± ± ± ±
0.5 0.05 0.05 0.05
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Fig. 4. Time evolution of foam volume for different mixtures (1 g/L) of aggregates/non-aggregated proteins (for protein aggregates with a Rh of 197 nm). Fig. 5. Time evolution of foam volume for mixtures (1 g/L) with 96% of protein aggregates for different aggregates sizes.
stable as the foam volume is, 1200 s after the end of bubbling, still higher than 30 ml. We observed that foamability and foam stability decrease with aggregates sizes. The time of foam formation was the same for foams made from protein aggregates with Rh 35 nm whereas for protein aggregates with Rh 71 nm, the time needed to form the foam was longer (120 s). For these foams the decrease of foam volume was faster than for those of non-aggregated proteins. For the large aggregates (117 and 197 nm), almost no foam was produced and the maximum foam volume could not be reached because the foams disappeared as fast as they were produced. In order to better understand the role of the aggregates in the formation and stabilization of foams, we made mixtures of nonaggregated proteins and protein aggregates with varying fraction of aggregates, ranging from 1 to 98%. The evolution of the foam volume for these mixtures is shown in Fig. 4 for protein aggregates of Rh 197 nm. The amount of protein aggregates in the mixture has a strong influence on the foam formation and stability. We observed two different types of behavior according to the amount of protein aggregates compared to foams obtained from non-aggregated proteins. For amounts ranging from 1 to 90%, no effect was observed on the foam formation (the time of foam formation was always about 80 s) but the stability of these foams was clearly improved compared with non-aggregated -lactoglobulin foams. Indeed, at the end of experiment, the foam volume of the foams made from mixtures with aggregates amounts ranging from 1 to 90% was higher than that of non-aggregated proteins foams. When the amount of protein aggregates was higher than 90%, it was more difficult to obtain the desired foam volume. Foam formation of foams made from mixtures with 92–96% of large protein aggregates was similar to foam made from non-aggregated proteins but the foam stability was strongly affected. The foam volume remaining 1200 s after the end of bubbling was lower than 20 mL. For mixtures with 98% of protein aggregates, the formation of the foam required a longer time and furthermore the foam destabilized very rapidly (only 5 ml of foam volume remained at the end of the experiment). For a given amount of protein aggregates especially at high amounts, foam stability depended on the aggregates size (Fig. 5). With a Rh of 35 and 71 nm, the sample containing 96% protein aggregates produced a more stable foam than non-aggregated proteins. For the same amount of protein aggregates, foams made from samples with Rh 117 and 197 nm were less stable. Moreover, the maximum amount of protein aggregates for which foams were more stable than that the ones obtained from non-aggregated proteins also depended of the aggregates size. For the large protein aggregates (117 and 197 nm), 90% is the maximum proportion for which the formed foams were more stable than that of non-aggregated proteins. At higher protein aggregate fractions (92–100%), foams were less stable (as shown in Fig. 4). For the small aggregates (35 and 71 nm), this maximum was 96%. Protein aggregates have not only an influence on the foam volume but also on the liquid volume and its evolution with time. In
our experiments, the total liquid volume incorporated in the foam increased when the aggregate size decreased (Table 2). However, it is important to consider not only the liquid volume but also the foam structure. For instance, the bubble size may be greatly affected by the aggregate size, leading to foams having the same liquid fraction but not the same structure. In order to study the effect of aggregate size on the drainage, we considered foams having the same foam volume, with equivalent stabilities and initial liquid fractions. This was possible by increasing the protein concentration. At 2 g/L of total protein concentration, the foams that were obtained had very close initial liquid volume whatever the aggregate size and contents. Moreover, the average bubble sizes estimated by optical microscopy were similar. For this reason, drainage experiments were performed from protein solutions with a concentration of 2 g/L. Fig. 6 shows the results obtained for time evolution of the normalized liquid fraction at electrode 1, i.e. at a fixed height of 6.5 cm above the base of the column. The liquid fraction is normalized by ε0 , defined as the liquid fraction at the end of foam formation when N2 bubbling has been stopped. This figure shows that the drainage rate was affected by the presence of aggregates. We observed a decrease of the drainage rate compared to non-aggregated -lactoglobulin whatever the amount of protein aggregates or the Rh . When the protein aggregates size increased, the drainage was slowed down. In the presence of 90% of protein aggregates (Fig. 6a), for the large protein aggregates (117 and 197 nm) the slope was twofold lower (slope −0.5) than that of foams stabilized by non-aggregated proteins (slope −1). For the small aggregates (35 and 71 nm), the slope had an intermediary value (slope −0.8). These results were still true even with 10% protein aggregates but the effect of the aggregate size seemed to be slightly more pronounced (Fig. 6b). 4. Discussion It has been shown in the literature [9,11,12] that the presence of protein aggregates can improve the foaming properties of a protein. Table 2 Liquid volume (in mL) incorporated in the foam measured just after foam formation as function of protein aggregates amount and size. The S.D. on the liquid volume values are of 0.2 mL % of aggregates 10
90
96
100
Non-aggregated -Ig
–
–
–
3.9
Protein aggregates size (nm) 35 71 117 197
4.2 4.7 3.8 4.0
4.1 3.8 2.4 2.8
3.3 2.7 1.6 1.8
3.6 1.4 No foam No foam
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Fig. 6. Time evolution of normalized liquid fraction for foams made from solutions at 2 g/L with, respectively: (a) 90% of protein aggregates and (b) 10% of protein aggregates.
Our results show that protein aggregates by themselves are not able to improve the foaming properties but they do participate in a better stabilization in the presence of non-aggregated proteins. The differences in the ability of protein aggregates to form foams can be mainly explained by differences in the adsorption kinetics, as observed in Fig. 2. The order in adsorption rate coincides with the protein aggregate size. Protein aggregates with a Rh of 197 nm cannot adsorb at the interface within the time scale of the foam experiment. Hence, no foam is formed. For protein aggregates with Rh 71 and 117 nm, the decrease of surface tension is the same. However, no foam is obtained from protein aggregates with a Rh of 117 nm, while it is possible to produce a foam with protein aggregates with a Rh of 71 nm. Drop tensiometer measurements at short times show that if foam formation is correlated with the rate at which the surface tension can be lowered by protein aggregates, this is not the only parameter that determines their ability to stabilize a foam. Foam formation is not only determined by the diffusion of protein to the interface but the protein also needs time to adsorb and anchor to the interface. Non-aggregated proteins adsorb to the interface more rapidly than protein aggregates because of their higher mobility. The observed differences in surface tension suggest that the mechanisms of adsorption and anchorage for protein aggregates at air/water interfaces are different from that of non-aggregated proteins. Large protein aggregates diffuse from the bulk to the interface and tend to adsorb. Because of their size, these aggregates occupy a high interfacial surface compared to their anchorage point at the surface, which can prevent other protein aggregates to adsorb close to them. As a consequence, we can assume that the interface is not completely covered by large protein aggregates. The second
stage of protein adsorption involves the unfolding of the molecule at the interface. Proteins within aggregates are likely to need a longer time to find a proper conformation to attach to the interface. This time may slow down the adsorption kinetics of protein aggregates compared with non-aggregated proteins. Zhu and Damodaran [19] suggested that non-aggregated proteins would adsorb more rapidly to an interface contributing to the foam formation. Aggregates would adsorb more slowly, thus contributing to the stability of an interfacial protein film that has already formed. Aggregates can increase the elasticity of the adsorbed layer, as shown by Davis and Foegeding [12]. An increase of the surface dilatational modulus has been related to a reduction of the foam destabilization mechanisms [20]. We note that surface tension measurements made at short times are not completely representative of protein adsorption in a foam. Saint-Jalmes et al. [13] have estimated the time of the formation of a bubble in a foam. This time is around 3 s, so protein adsorption should take place during this very short time in order to produce a stable bubble. This time is small compared to the anchoring time of proteins. The consequence is that after formation, the liquid–gas interfaces of the bubbles are probably widely uncovered by stabilizing proteins. These authors suggest that in situ surface tension measurements (that means in a real foam film made after the adsorption of the aggregates) should give a more relevant information. Moreover, while a model interface such as the one used in the drop tensiometer can be observed for hours or days before equilibrium is reached, some foam proteins destabilize in a few minutes after bubbling. This indicates that the foaming properties of proteins are not measured under the same conditions as the drop tensiometer measurements.
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Foaming experiments show that a given amount of nonaggregated proteins is required in order to form a stable foam. When only protein aggregates are present, no foams are obtained (for the large aggregates) or the foams are less stable than non-aggregated proteins (for the small aggregates). The minimum quantity of nonaggregated proteins needed to stabilize a foam depends on the aggregate size. This required quantity decreases with the aggregate size. This fact can be related to the more efficient adsorption of small aggregates (35 and 71 nm) observed in the drop tensiometer measurements and to the capacity of these aggregates to partly stabilize foams. Due to their size, the steric effect is certainly less important and protein aggregates can cover the interface more easily. If the amount of non-aggregated proteins are sufficient to form a stable foam, protein aggregates will improve the foaming properties. This result has already been observed by Schmitt et al. [9,10]. It is interesting to note that even a very low amount of protein aggregates (1%) is enough to improve the foaming properties compared with non-aggregated proteins. No change of the surface tension was observed by drop tensiometer measurements for samples with 1% of protein aggregates compared to the non-aggregated proteins. For amounts of protein aggregates ranging from 1 to 90%, non-aggregated proteins are sufficient to cover the bubble in the drop tensiometer and then control the kinetic adsorption of the mixtures. This confirms that the ability of aggregates to decrease the surface tension is not the only parameter that influences their foaming properties. We observed a decrease of the liquid fraction in the foam depending on the quantity of aggregates and the aggregate size (see Table 2). As large protein aggregates alone are not able to produce foams, it seems logical that when the amount of aggregates increases, the resulting foam contains less liquid. Indeed, the increase of protein aggregates led to foam constituted of large bubbles, filling more volume for the same quantity of liquid. Table 2 then is difficult to interpret because foams made from nonaggregated proteins and mixtures with various fractions of protein aggregates do not have the same macroscopic features (bubbles size and number). In order to check if the improvement of the foaming properties was not due to a concentration effect in the foam, we determined the total protein concentration and the respective parts of nonaggregated and aggregated proteins in the foams by GFC and UV absorbance. The liquid volume remaining in the cuvette sample was analyzed and by difference, the non-aggregated protein and aggregate concentrations in the foams could be obtained. From these measurements, we conclude that whatever the foam (in terms of size and quantity of protein aggregates), the presence of protein aggregates did not change the total protein concentration in the foam. This suggests that the improvement of the foam stability by protein aggregates is not due to an increase of the total protein concentration in the foam. Moreover, the same ratio of non-aggregated proteins/protein aggregates was found in the initial solution before bubbling and in the foam just after bubbling. The presence of aggregates does have an influence on the drainage, as shown by comparing foams with similar macroscopic texture and stability (foams obtained from total protein concentration at 2 g/L). When comparing foams having the same physical properties, we observed that the drainage is slowed down when the size of protein aggregates increases. We suggest two different possible and coexisting mechanisms for foam stabilization by protein aggregates. Firstly, the aggregates may form a viscoelastic layer in the thin films between bubbles, leading to a gel-like network and in this way contributing to the foam stabilization. This effect has been shown by Schmitt et al.
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[10] for WPI aggregates. Moreover, Saint-Jalmes et al. [13] suggested that even if the interface is poorly covered, stable foams could still be produced by the confinement of protein aggregates in the foam films. In this case, a percolation process may then lead to more rigid films which could avoid any flow or drainage and thus increase the foam stability. Protein aggregation can also lead to cross-linking between the interfaces of a thin film, giving rise to stable films [21]. Due to their size and the lack of interfacial properties observed on drop tensiometer, it is also reasonable to suggest that large protein aggregates remains in the liquid phase and may be located either in the thin films or in the Plateau borders. In that case, protein aggregates may play a role as cork which could prevent foam drainage. This kind of stabilizing mechanism has already been observed for foams stabilized by non-adsorbing nanoparticles [22,23]. We can consider protein aggregates as nanoparticles having specific surface properties. Nanoparticles create an effective steric clutter, affecting the film between bubbles. This will avoid coalescence and by this way cause the retardation of the drainage. This could partly explain the stability of foams made from samples with very low amounts of large protein aggregates. In that case, it seems difficult that aggregates form a gel-like network in the films. However, we observed that these samples produced foams that are more stable than foams from non-aggregated proteins alone. We can then imagine that for small amounts of large aggregates, the stabilization by a cork effect will be predominant. The rate of drainage should be related to the aggregate size. Indeed, we observed that the rate of drainage was affected by the aggregate size more strongly for small amounts of aggregates than for large amounts (see Fig. 6). 5. Conclusion We have reported the effect of protein aggregates on the foaming properties of non-aggregated -lactoglobulin. We have shown that the amount and the size of aggregates have an influence on foam formation and stability. Non-aggregated proteins appear to be necessary to produce stable foams. Protein foams containing aggregates can be stabilized either by the formation of viscoelastic layers in the thin films or by the confinement of protein aggregates in the Plateau borders, both mechanisms leading to a decrease of the drainage rate. Thin liquid film experiments and measurements of the viscoelasticity of adsorbed layers will be done in order to determine the behavior of protein aggregates in films. In particular, it will be interesting to check if protein aggregates can interact with the interface and under which conditions. Acknowledgements We thank Dr. Dominique Durand and Dr. Taco Nicolai for help with the static light scattering measurements performed in Polymers Colloids and Interfaces Laboratory, Université du Maine, Le Mans, France. We also thank Hélène Rogniaux, and Audrey Geairon for technical support during the mass spectrometry analyses conducted by the platform “Biopolymers-Interaction-Structural Biology” located at the INRA Center of Nantes (INRA Research Unit 1268). References [1] D. Weaire, S. Hutzler, The Physics of Foams, Clarendon Press, Oxford, 1999. [2] B.S. Murray, Stabilization of bubbles and foams, Current Opinion in Colloid & Interface Science 12 (2007) 232–241. [3] E.A. Foegeding, P.J. Luck, J.P. Davis, Factors determining the physical properties of protein foams, Food Hydrocolloids 20 (2006) 284–292.
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[14] M. Pouzot, D. Durand, T. Nicolai, Influence of the ionic strength on the structure of heat-set globular protein gels at pH 7, Macromolecules 37 (2004) 8703–8708. [15] N. Mahmoudi, S. Mehalebi, T. Nicolai, D. Durand, A. Riaublanc, Light-scattering study of the structure of aggregates and gels formed by heat-denatured whey protein isolate and -lactoglobulin at neutral pH, Journal of Agricultural and Food Chemistry 55 (2007) 3104–3111. [16] C. Le Bon, T. Nicolai, D. Durand, Growth and structure of aggregates of heatinduced -lactoglobulin, International Journal of Food Science and Technology 34 (1999) 451–465. [17] P. Maillart, B. Ribadeau-Dumas, Preparation of -lactoglobulin and lactoglobulin-free proteins from whey retentate by NaCl salting out at low pH, Journal of Food Science 53 (1988) 743–745. [18] J. Benjamins, A. Cagna, E.H. Lucassen-Reynders, Viscoelastic properties of triacylglycerol/water interfaces covered by proteins, Colloids and Surfaces A: Physicochemical and Engineering Aspects 114 (1996) 245–254. [19] H. Zhu, S. Damodaran, Heat-induced conformational changes in whey protein isolate and its relation to foaming properties, Journal of Agricultural and Food Chemistry 42 (1994) 846–855. [20] A. Prins, M.A. Bos, F.J.G. Boerboom, H. Van Kalsbeek, Relation between surface rheology and foaming behavior of aqueous protein solutions, in: D. Mobius, R. Miller (Eds.), Proteins at Liquid Interfaces, Elsevier Science, Amsterdam, 1998, pp. 221–266. [21] T.D. Dimitrova, F. Leal-Calderon, T.D. Gurkov, B. Campbell, Surfaces forces in model oil-in-water emulsions stabilized by proteins, Advances in Colloid and Interface Science 108 (2004) 73–86. [22] B.S. Murray, R. Ettelaie, Foam stability: proteins and nanoparticles, Current Opinion in Colloid & Interface Science 9 (2004) 314–320. [23] T.N. Hunter, R.J. Pugh, G.V. franks, G.J. Jameson, The role of particles in stabilizing foams and emulsions, Advances in Colloid and Interface Science 137 (2008) 57–81.
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Colloids and Surfaces A: Physicochemical and Engineering Aspects journal homepage: www.elsevier.com/locate/colsurfa
Effect of protein aggregates on foaming properties of -lactoglobulin Bénédicte Rullier, Bruno Novales ∗ , Monique A.V. Axelos Institut National de la Recherche Agronomique, UR1268 Biopolymères, Interactions, Assemblages, INRA, Equipe Interfaces Systèmes Dispersés, Rue de la Géraudière, 44316 Nantes Cedex 03, France
a r t i c l e
i n f o
Article history: Received 3 June 2008 Received in revised form 16 July 2008 Accepted 19 July 2008 Available online 30 July 2008 Keywords: Foam Protein aggregates Interfacial properties
a b s t r a c t Our paper aims at determining the respective part of protein aggregates and non-aggregated proteins in the foam formation and stability of -lactoglobulin. We report results on fractal aggregates formed at neutral pH and strong ionic strength (aggregates size from 30 to 190 nm). Pure aggregates and mixtures of non-aggregated/aggregated proteins at varying ratios were used. The capacity of aggregates to form and stabilize foams has been studied in relation with their ability to absorb at air/water interfaces. Our results show that protein aggregates are not able by themselves to improve the foaming properties but participate to a better foam stabilization in the presence of non-aggregated proteins. Non-aggregated proteins appear to be necessary to produce stable foams. We have shown that the amount and the size of aggregates had an influence on the drainage rate. © 2008 Elsevier B.V. All rights reserved.
1. Introduction Aqueous foams are dispersions of gas bubbles in a liquid stabilized by amphiphilic molecules [1]. In the food industry, foams are stabilized mainly by proteins. A large number of papers have been devoted to the characterization of the foaming behavior of proteins under a variety of conditions [2,3]. Surfactants rapidly decrease the surface tension to low values, while proteins slowly but strongly adsorb to the air/water interface. Even if the decrease of surface tension is less important for proteins, it has been demonstrated to be correlated to their foaming ability [4–6]. In particular, it has been shown that flexible proteins with high surface activity adsorb more rapidly to the interface than compact globular proteins [7]. When adsorbed at the interface, proteins create high viscoelastic layers with high surface shear viscosities. Martin et al. [7] have been demonstrated that under certain conditions, the adsorbed protein film can form a resistant and cohesive interfacial network which enhances foam stability. Most of the studies cited have been carried out on nonaggregated proteins. However, during many food processings, proteins undergo thermomechanical treatments that often lead to their aggregation in self-assembled structures [8–10]. As protein aggregates are complex structures whose size may be as large as a few hundred nanometers, it can be expected that their behavior is different from non-aggregated proteins. Because of their size, the ability of aggregates to diffuse to the interface is reduced and this
∗ Corresponding author. Tel.: +33 240675198; fax: +33 240675084. E-mail address: [email protected] (B. Novales). 0927-7757/$ – see front matter © 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.colsurfa.2008.07.040
should lead to lower foaming properties. Moreover, protein–protein interactions within aggregates can be of a different nature (electrostatic, hydrophobic, etc.), leading to different kinds of cohesiveness. This could affect the mobility of proteins and their ability to spread at the interface and influence their capacity to stabilize air/water interfaces needed to produce a foam. However, it has been shown that the presence of some aggregates increases the foaming stability [9,11,12]. The mechanism by which aggregates contribute to foam stabilization is not completely elucidated. Different hypotheses have been proposed: either the protein aggregates can adsorb at the interface or they cannot. If they do, aggregates increase the viscoelasticity of the interface thus providing a better foam stabilization [12]. On the other hand, if aggregates do not adsorb to the interface, they remain in the aqueous phase. In this case, aggregates can become confined into foam films and can undergo a percolation process leading to the formation of a gel-like network. They can also prevent foam destabilization by reducing drainage acting as cork in the Plateau borders [13]. In industrial processes, only a given fraction of the proteins is aggregated and the part of non-aggregated proteins may considerably influence the foaming properties. Therefore, it is often difficult to determine the intrinsic role of aggregates in these systems. The aim of our paper is to determine the respective part of protein aggregates and of non-aggregated proteins played in the foam formation and foam stability of protein solutions. For the present work -lactoglobulin has been chosen because its aggregation properties in function of experimental parameters such as temperature, pH, ionic strength and protein concentration have been studied in considerable detail [14–16]. The size and the shape of -lactoglobulin aggregates can be easily varied by the choice of the experimen-
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tal conditions, leading to fibrillar or fractal aggregates with sizes ranging from a few nanometers to hundreds of nanometers. In this paper, we report results on fractal aggregates exclusively formed by heating at 80 ◦ C during 24 h at neutral pH and 0.1 M of ionic strength. The protein concentration was varied in order to modulate the size of the aggregates. The capacity of the aggregates to form and stabilize foams has been studied in relation with their ability to absorb at air/water interfaces.
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The amount of non-aggregated protein after the UF–C step was determined by gel filtration chromatography (GFC) under the following conditions: column TSK PW 6000 equilibrated with an elution solution composed of Trizma NH2 C(CH2 OH)3 0.05 M; NaCl 0.1 M at pH 7. The eluted protein was detected using UV adsorption at 280 nm and the amount of residual non-aggregated protein was calculated as percent area relative to the area of unheated protein. 2.3. Surface tension measurements
2. Materials and methods 2.1. ˇ-Lactoglobulin -Lactoglobulin was purified from whey protein isolate (Prolacta 90 batch no. 273 supplied by Lactalis Industrie, Laval, France) according to the method of Maillart and Ribadeau-Dumas [17]. After lyophilisation, -lactoglobulin was analyzed by mass spectrometry. Each sample of -lactoglobulin was constituted of 55% of -IgA, 35% of -IgB and 10% of -IgA lactosylated. The average mass was determined at 18,385 kDa. -Lactoglobulin powder was dissolved in milliQ water at pH 7 containing 3 mM sodium azide to prevent from bacterial contamination. -Lactoglobulin solution was dialyzed and filtered with a 0.22 m membrane. Protein solutions were then dissolved in a 0.1 M sodium chloride (NaCl) solution at pH 7. Protein concentrations were determined through the measurement of UV adsorption at 278 nm, taken an extinction coefficient of 0.96 L g−1 cm [16]. 2.2. Formation and characterization of protein aggregates -Lactoglobulin aggregates were formed upon heat-induced denaturation of proteins at different concentrations at a temperature of 80 ◦ C during 24 h. The samples were cooled in a bath at room temperature. In this paper, aggregates were formed at pH 7 and 0.1 M NaCl ionic strength at four concentrations: 1, 2, 8 and 10 g/L. Aggregate sizes were determined by static and dynamic light scattering. Static measurements were made using an ALV-5000 multi-bit multi-tau correlator and a Spectra Physics solid-state laser operating with polarized light with wavelength = 532 nm. The range of scattering wave vectors covered was from 3.0 × 10−3 to 3.5 × 10−2 nm−1 (q = 4ns sin(/2)/, with the angle of observation and ns the refractive index of the solution). Dynamic light scattering was performed using a Nanosizer ZS (Malvern Instruments, UK) in order to determine the hydrodynamic radius of the aggregates. The instrument was used in the backscattering configuration where detection is done at a scattering angle of 173◦ . Dilute solutions were measured in a 1-cm path-length spectroscopic plastic cell at 20 ◦ C. The hydrodynamic radius was measured in triplicate. Each measurement corresponded to three autocorrelation functions recorded during 90s. As the size of our aggregates were close to the range of reliability of the Malvern Nanosizer (given by the manufacturer), data obtained from this apparatus were compared to results obtained by static light scattering for some samples in order to check the reliability of our measurements. In order to obtain solutions containing exclusively protein aggregates, the non-aggregated proteins remaining in the solution were separated out by an ultrafiltration–centrifugation (UF–C) using an Amicon membrane Ultra 30 kDa. This step was performed during 30 min at 4000 rpm and 4 ◦ C for the larger aggregates, and during 4 h for the smaller ones. The retentate was redispersed in a 0.1 M NaCl solution and the concentration was verified by the measurement of UV absorbance at 278 nm.
An automated drop tensiometer (IT-Concept, Longessaigne, France) was used to measure the interfacial tension between liquid and air, as described in detail by Benjamins et al. [18]. All experiments were performed in the rising bubble configuration. A 6 L air bubbles was formed upward at the tip of a U-shaped stainless steel needle immersed in a cuvette filled with the protein solution, under gentle stirring, both of which are temperature controlled at 20 ◦ C. The dynamic tension was followed in time (120 s in our case) using axisymmetric drop-shape analysis. Image acquisition and regression of the interfacial tension were performed with Windrop software by fitting the Laplace equation to the drop shape. 2.4. Foam experiments Foaming experiments were conducted on a “Foamscan” apparatus developed by IT Concept (Longessaigne, France). With this instrument, the foam formation, the stability and the drainage of liquid from the foam can be determined by conductimetric and optical measurements. After calibration of the conductimetric electrodes, foams were produced from 12 mL of the solution at protein concentration 1 g/L by injection of gaseous N2 through a porous glass filter. The flow rate was fixed at 35 mL/min. All foams were allowed to reach a final volume of 45 mL after which gas-flow was stopped and the evolution of the foam was analyzed. The foamability corresponds to the time needed to reach 45 mL of foam volume. After foam formation, in a timespan of 3600 s, several parameters are automatically recorded by the “Foamscan” analyzing software. The volume of the foam is determined with a CCD camera (Sony Exwave HAD). A pair of electrodes at the bottom of the column is used to measure the quantity of liquid that is not in the foam, while the volume of the liquid in the foam is measured by conductimetry using three pairs of electrodes located along the glass column. The liquid drainage of the foam is followed via conductivity measurements at different heights in the foam column. In order to get information about the composition of the protein solutions and foams, the drained liquid was quantified using the GFC method previously described in detail. These measurements were made at two different times: firstly just after the foam formation when the desired foam volume is reached and secondly 1200 s after the end of bubbling. For long-time drainage experiments (3600 s), in order to check that the drainage was not affected by other destabilization mechanisms (such as coarsening), some foams were produced by injecting N2 through a perfluorohexane (C6 F14 ) solution. We observed that at the electrode 1 (where the liquid fraction is measured), the drainage curves had similar profiles with or without C6 F14 . 3. Results 3.1. Characterization of protein aggregates From static light scattering measurements, the dependence of the scattered intensity Ir (q) on the scattering wave vector (q) by the aggregates at different protein concentrations can be obtained.
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Fig. 2. Dynamic surface tension measurements of non-aggregated proteins and protein aggregates at C = 0.05 g/L: (䊉) non-aggregated protein; aggregates () 35 nm; () 71 nm; (×) 117 nm; ()197 nm.
the rest of this paper, when referring to protein aggregate samples their hydrodynamic radius measured after UF–C is meant. Fig. 1. Master curves of normalized scattering wave vector (q) as function of the normalized intensity of scattered light (Ir ) for dilute solutions of -lactoglobulin aggregates.
From the initial q-dependence the z-average radius of gyration (Rgz ) of the aggregates can be extracted. After dividing Ir (q) by the weight average molar mass Mw and plotting the result as a function of qRgz , a master curve is obtained (Fig. 1) which means that aggregates are self-similar. At high qRgz -values, we observe a power law dependence of Ir (q) ∞ q−df . The exponent df represents the socalled fractal dimension of the aggregates [16]. In our case, df is close to 2 which corresponds to typical fractal aggregates. From the weight average molar mass Mw determined from the curves, it is possible to determine the average number of proteins within aggregates. The measured number ranges from a few hundred proteins for the small aggregates to around 5000 proteins for the larger ones. From dynamic light scattering the size distribution of nonaggregated proteins as well as of protein aggregates was obtained. Non-aggregated -lactoglobulin has a hydrodynamic radius of 3 nm and the weight molar mass Mw was determine at 33 kg/mol, which is coherent with data in the literature [15]. The average hydrodynamic radius (Rh ) of protein aggregates are reported in Table 1. The aggregate size increases with protein concentration with Rh ranging from 35 nm at 1 g/L to 197 nm at 10 g/L. By contrast, the amount of non-aggregated protein, determined by GFC, increases when the protein concentration decreases. When the heat treatment is stopped after 24 h, the equilibrium has not yet been reached and as the aggregation process depends on time and protein concentration, this explains the observed variations of aggregate size and aggregates/non-aggregated protein ratios. In order to obtain exclusively protein aggregates, nonaggregated proteins were separated from the aggregates by UF–C. After the UF–C step, the aggregate size was checked by dynamic light scattering, and the amount of non-aggregated proteins was determined by GFC to be close to 0.5% for large aggregates (117 and 197 nm) and close to 1% for small aggregates (35 and 71 nm). For
3.2. Surface tension As foam formation occurs in a few seconds, it is more pertinent to study the adsorption kinetics at short times when attempting to explain the ability of proteins to adsorb at air/water interfaces. The surface tension of both non-aggregated proteins and protein aggregates measured during 120 s is shown in Fig. 2. For non-aggregated -lactoglobulin, we observed the fastest decrease of the surface tension, with a value lower than 60 mN/m after 120 s. For protein aggregates, the adsorption kinetic is slower. The adsorption rate of the aggregates decreases with their size. Three groups of adsorption rate behavior can be distinguish: (i) for the largest protein aggregates no decrease of the surface tension was observed; (ii) intermediate size aggregates with Rh 71 and 117 nm; and (iii) small protein aggregates (Rh 35 nm) which, during the first 10 s, present a decrease of surface tension that is similar to that of non-aggregated proteins. 3.3. Foaming properties The time evolution of the foam volume for non-aggregated proteins and for protein aggregates is shown in Fig. 3. A foam made from non-aggregated proteins is formed in 82 s. Such a foam is relatively
Fig. 3. Time evolution of foam volume for non-aggregated proteins and proteins aggregates solutions (1 g/L).
Table 1 -Lactoglobulin aggregates hydrodynamic radius and fractions of non-aggregated proteins measured before and after UF–C step as function of protein concentration Protein concentration (g/L)
Before UF–C Rh (nm)
1 2 8 10
34 66 111 196
± ± ± ±
6 1 1 5
After UF–C % of non-aggregated proteins 20.4 10.4 10.6 3.7
± ± ± ±
10.0 2.0 2.0 0.6
Rh (nm) 35 71 117 197
± ± ± ±
1 1 1 10
% of non-aggregated proteins 1.0 0.80 0.80 0.50
± ± ± ±
0.5 0.05 0.05 0.05
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Fig. 4. Time evolution of foam volume for different mixtures (1 g/L) of aggregates/non-aggregated proteins (for protein aggregates with a Rh of 197 nm). Fig. 5. Time evolution of foam volume for mixtures (1 g/L) with 96% of protein aggregates for different aggregates sizes.
stable as the foam volume is, 1200 s after the end of bubbling, still higher than 30 ml. We observed that foamability and foam stability decrease with aggregates sizes. The time of foam formation was the same for foams made from protein aggregates with Rh 35 nm whereas for protein aggregates with Rh 71 nm, the time needed to form the foam was longer (120 s). For these foams the decrease of foam volume was faster than for those of non-aggregated proteins. For the large aggregates (117 and 197 nm), almost no foam was produced and the maximum foam volume could not be reached because the foams disappeared as fast as they were produced. In order to better understand the role of the aggregates in the formation and stabilization of foams, we made mixtures of nonaggregated proteins and protein aggregates with varying fraction of aggregates, ranging from 1 to 98%. The evolution of the foam volume for these mixtures is shown in Fig. 4 for protein aggregates of Rh 197 nm. The amount of protein aggregates in the mixture has a strong influence on the foam formation and stability. We observed two different types of behavior according to the amount of protein aggregates compared to foams obtained from non-aggregated proteins. For amounts ranging from 1 to 90%, no effect was observed on the foam formation (the time of foam formation was always about 80 s) but the stability of these foams was clearly improved compared with non-aggregated -lactoglobulin foams. Indeed, at the end of experiment, the foam volume of the foams made from mixtures with aggregates amounts ranging from 1 to 90% was higher than that of non-aggregated proteins foams. When the amount of protein aggregates was higher than 90%, it was more difficult to obtain the desired foam volume. Foam formation of foams made from mixtures with 92–96% of large protein aggregates was similar to foam made from non-aggregated proteins but the foam stability was strongly affected. The foam volume remaining 1200 s after the end of bubbling was lower than 20 mL. For mixtures with 98% of protein aggregates, the formation of the foam required a longer time and furthermore the foam destabilized very rapidly (only 5 ml of foam volume remained at the end of the experiment). For a given amount of protein aggregates especially at high amounts, foam stability depended on the aggregates size (Fig. 5). With a Rh of 35 and 71 nm, the sample containing 96% protein aggregates produced a more stable foam than non-aggregated proteins. For the same amount of protein aggregates, foams made from samples with Rh 117 and 197 nm were less stable. Moreover, the maximum amount of protein aggregates for which foams were more stable than that the ones obtained from non-aggregated proteins also depended of the aggregates size. For the large protein aggregates (117 and 197 nm), 90% is the maximum proportion for which the formed foams were more stable than that of non-aggregated proteins. At higher protein aggregate fractions (92–100%), foams were less stable (as shown in Fig. 4). For the small aggregates (35 and 71 nm), this maximum was 96%. Protein aggregates have not only an influence on the foam volume but also on the liquid volume and its evolution with time. In
our experiments, the total liquid volume incorporated in the foam increased when the aggregate size decreased (Table 2). However, it is important to consider not only the liquid volume but also the foam structure. For instance, the bubble size may be greatly affected by the aggregate size, leading to foams having the same liquid fraction but not the same structure. In order to study the effect of aggregate size on the drainage, we considered foams having the same foam volume, with equivalent stabilities and initial liquid fractions. This was possible by increasing the protein concentration. At 2 g/L of total protein concentration, the foams that were obtained had very close initial liquid volume whatever the aggregate size and contents. Moreover, the average bubble sizes estimated by optical microscopy were similar. For this reason, drainage experiments were performed from protein solutions with a concentration of 2 g/L. Fig. 6 shows the results obtained for time evolution of the normalized liquid fraction at electrode 1, i.e. at a fixed height of 6.5 cm above the base of the column. The liquid fraction is normalized by ε0 , defined as the liquid fraction at the end of foam formation when N2 bubbling has been stopped. This figure shows that the drainage rate was affected by the presence of aggregates. We observed a decrease of the drainage rate compared to non-aggregated -lactoglobulin whatever the amount of protein aggregates or the Rh . When the protein aggregates size increased, the drainage was slowed down. In the presence of 90% of protein aggregates (Fig. 6a), for the large protein aggregates (117 and 197 nm) the slope was twofold lower (slope −0.5) than that of foams stabilized by non-aggregated proteins (slope −1). For the small aggregates (35 and 71 nm), the slope had an intermediary value (slope −0.8). These results were still true even with 10% protein aggregates but the effect of the aggregate size seemed to be slightly more pronounced (Fig. 6b). 4. Discussion It has been shown in the literature [9,11,12] that the presence of protein aggregates can improve the foaming properties of a protein. Table 2 Liquid volume (in mL) incorporated in the foam measured just after foam formation as function of protein aggregates amount and size. The S.D. on the liquid volume values are of 0.2 mL % of aggregates 10
90
96
100
Non-aggregated -Ig
–
–
–
3.9
Protein aggregates size (nm) 35 71 117 197
4.2 4.7 3.8 4.0
4.1 3.8 2.4 2.8
3.3 2.7 1.6 1.8
3.6 1.4 No foam No foam
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Fig. 6. Time evolution of normalized liquid fraction for foams made from solutions at 2 g/L with, respectively: (a) 90% of protein aggregates and (b) 10% of protein aggregates.
Our results show that protein aggregates by themselves are not able to improve the foaming properties but they do participate in a better stabilization in the presence of non-aggregated proteins. The differences in the ability of protein aggregates to form foams can be mainly explained by differences in the adsorption kinetics, as observed in Fig. 2. The order in adsorption rate coincides with the protein aggregate size. Protein aggregates with a Rh of 197 nm cannot adsorb at the interface within the time scale of the foam experiment. Hence, no foam is formed. For protein aggregates with Rh 71 and 117 nm, the decrease of surface tension is the same. However, no foam is obtained from protein aggregates with a Rh of 117 nm, while it is possible to produce a foam with protein aggregates with a Rh of 71 nm. Drop tensiometer measurements at short times show that if foam formation is correlated with the rate at which the surface tension can be lowered by protein aggregates, this is not the only parameter that determines their ability to stabilize a foam. Foam formation is not only determined by the diffusion of protein to the interface but the protein also needs time to adsorb and anchor to the interface. Non-aggregated proteins adsorb to the interface more rapidly than protein aggregates because of their higher mobility. The observed differences in surface tension suggest that the mechanisms of adsorption and anchorage for protein aggregates at air/water interfaces are different from that of non-aggregated proteins. Large protein aggregates diffuse from the bulk to the interface and tend to adsorb. Because of their size, these aggregates occupy a high interfacial surface compared to their anchorage point at the surface, which can prevent other protein aggregates to adsorb close to them. As a consequence, we can assume that the interface is not completely covered by large protein aggregates. The second
stage of protein adsorption involves the unfolding of the molecule at the interface. Proteins within aggregates are likely to need a longer time to find a proper conformation to attach to the interface. This time may slow down the adsorption kinetics of protein aggregates compared with non-aggregated proteins. Zhu and Damodaran [19] suggested that non-aggregated proteins would adsorb more rapidly to an interface contributing to the foam formation. Aggregates would adsorb more slowly, thus contributing to the stability of an interfacial protein film that has already formed. Aggregates can increase the elasticity of the adsorbed layer, as shown by Davis and Foegeding [12]. An increase of the surface dilatational modulus has been related to a reduction of the foam destabilization mechanisms [20]. We note that surface tension measurements made at short times are not completely representative of protein adsorption in a foam. Saint-Jalmes et al. [13] have estimated the time of the formation of a bubble in a foam. This time is around 3 s, so protein adsorption should take place during this very short time in order to produce a stable bubble. This time is small compared to the anchoring time of proteins. The consequence is that after formation, the liquid–gas interfaces of the bubbles are probably widely uncovered by stabilizing proteins. These authors suggest that in situ surface tension measurements (that means in a real foam film made after the adsorption of the aggregates) should give a more relevant information. Moreover, while a model interface such as the one used in the drop tensiometer can be observed for hours or days before equilibrium is reached, some foam proteins destabilize in a few minutes after bubbling. This indicates that the foaming properties of proteins are not measured under the same conditions as the drop tensiometer measurements.
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Foaming experiments show that a given amount of nonaggregated proteins is required in order to form a stable foam. When only protein aggregates are present, no foams are obtained (for the large aggregates) or the foams are less stable than non-aggregated proteins (for the small aggregates). The minimum quantity of nonaggregated proteins needed to stabilize a foam depends on the aggregate size. This required quantity decreases with the aggregate size. This fact can be related to the more efficient adsorption of small aggregates (35 and 71 nm) observed in the drop tensiometer measurements and to the capacity of these aggregates to partly stabilize foams. Due to their size, the steric effect is certainly less important and protein aggregates can cover the interface more easily. If the amount of non-aggregated proteins are sufficient to form a stable foam, protein aggregates will improve the foaming properties. This result has already been observed by Schmitt et al. [9,10]. It is interesting to note that even a very low amount of protein aggregates (1%) is enough to improve the foaming properties compared with non-aggregated proteins. No change of the surface tension was observed by drop tensiometer measurements for samples with 1% of protein aggregates compared to the non-aggregated proteins. For amounts of protein aggregates ranging from 1 to 90%, non-aggregated proteins are sufficient to cover the bubble in the drop tensiometer and then control the kinetic adsorption of the mixtures. This confirms that the ability of aggregates to decrease the surface tension is not the only parameter that influences their foaming properties. We observed a decrease of the liquid fraction in the foam depending on the quantity of aggregates and the aggregate size (see Table 2). As large protein aggregates alone are not able to produce foams, it seems logical that when the amount of aggregates increases, the resulting foam contains less liquid. Indeed, the increase of protein aggregates led to foam constituted of large bubbles, filling more volume for the same quantity of liquid. Table 2 then is difficult to interpret because foams made from nonaggregated proteins and mixtures with various fractions of protein aggregates do not have the same macroscopic features (bubbles size and number). In order to check if the improvement of the foaming properties was not due to a concentration effect in the foam, we determined the total protein concentration and the respective parts of nonaggregated and aggregated proteins in the foams by GFC and UV absorbance. The liquid volume remaining in the cuvette sample was analyzed and by difference, the non-aggregated protein and aggregate concentrations in the foams could be obtained. From these measurements, we conclude that whatever the foam (in terms of size and quantity of protein aggregates), the presence of protein aggregates did not change the total protein concentration in the foam. This suggests that the improvement of the foam stability by protein aggregates is not due to an increase of the total protein concentration in the foam. Moreover, the same ratio of non-aggregated proteins/protein aggregates was found in the initial solution before bubbling and in the foam just after bubbling. The presence of aggregates does have an influence on the drainage, as shown by comparing foams with similar macroscopic texture and stability (foams obtained from total protein concentration at 2 g/L). When comparing foams having the same physical properties, we observed that the drainage is slowed down when the size of protein aggregates increases. We suggest two different possible and coexisting mechanisms for foam stabilization by protein aggregates. Firstly, the aggregates may form a viscoelastic layer in the thin films between bubbles, leading to a gel-like network and in this way contributing to the foam stabilization. This effect has been shown by Schmitt et al.
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[10] for WPI aggregates. Moreover, Saint-Jalmes et al. [13] suggested that even if the interface is poorly covered, stable foams could still be produced by the confinement of protein aggregates in the foam films. In this case, a percolation process may then lead to more rigid films which could avoid any flow or drainage and thus increase the foam stability. Protein aggregation can also lead to cross-linking between the interfaces of a thin film, giving rise to stable films [21]. Due to their size and the lack of interfacial properties observed on drop tensiometer, it is also reasonable to suggest that large protein aggregates remains in the liquid phase and may be located either in the thin films or in the Plateau borders. In that case, protein aggregates may play a role as cork which could prevent foam drainage. This kind of stabilizing mechanism has already been observed for foams stabilized by non-adsorbing nanoparticles [22,23]. We can consider protein aggregates as nanoparticles having specific surface properties. Nanoparticles create an effective steric clutter, affecting the film between bubbles. This will avoid coalescence and by this way cause the retardation of the drainage. This could partly explain the stability of foams made from samples with very low amounts of large protein aggregates. In that case, it seems difficult that aggregates form a gel-like network in the films. However, we observed that these samples produced foams that are more stable than foams from non-aggregated proteins alone. We can then imagine that for small amounts of large aggregates, the stabilization by a cork effect will be predominant. The rate of drainage should be related to the aggregate size. Indeed, we observed that the rate of drainage was affected by the aggregate size more strongly for small amounts of aggregates than for large amounts (see Fig. 6). 5. Conclusion We have reported the effect of protein aggregates on the foaming properties of non-aggregated -lactoglobulin. We have shown that the amount and the size of aggregates have an influence on foam formation and stability. Non-aggregated proteins appear to be necessary to produce stable foams. Protein foams containing aggregates can be stabilized either by the formation of viscoelastic layers in the thin films or by the confinement of protein aggregates in the Plateau borders, both mechanisms leading to a decrease of the drainage rate. Thin liquid film experiments and measurements of the viscoelasticity of adsorbed layers will be done in order to determine the behavior of protein aggregates in films. In particular, it will be interesting to check if protein aggregates can interact with the interface and under which conditions. Acknowledgements We thank Dr. Dominique Durand and Dr. Taco Nicolai for help with the static light scattering measurements performed in Polymers Colloids and Interfaces Laboratory, Université du Maine, Le Mans, France. We also thank Hélène Rogniaux, and Audrey Geairon for technical support during the mass spectrometry analyses conducted by the platform “Biopolymers-Interaction-Structural Biology” located at the INRA Center of Nantes (INRA Research Unit 1268). References [1] D. Weaire, S. Hutzler, The Physics of Foams, Clarendon Press, Oxford, 1999. [2] B.S. Murray, Stabilization of bubbles and foams, Current Opinion in Colloid & Interface Science 12 (2007) 232–241. [3] E.A. Foegeding, P.J. Luck, J.P. Davis, Factors determining the physical properties of protein foams, Food Hydrocolloids 20 (2006) 284–292.
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