Journal of Dairy Science
Volume 92, Issue 4 , Pages 1387-1397, April 2009

Functional properties of whey proteins affected by heat treatment and hydrodynamic high-pressure shearing

School of Biomedical and Health Sciences, Faculty of Health, Engineering and Science, Victoria University, PO Box 14428, Melbourne, Victoria 8001 Australia

Received 7 October 2008; accepted 13 November 2008.

Article Outline

Abstract 

Two batches of native whey proteins (WP) were subjected to microfluidization or heat denaturation accompanied by microfluidization, followed by spray drying. Powders were assessed for their solubility, heat stability, coagulation time, and emulsifying and foaming properties. Effects of denaturation and shearing were examined by particle size analysis, differential scanning calorimetry, reducing and nonreducing sodium dodecyl sulfate-PAGE, and size exclusion-HPLC. Heat treatment significantly decreased solubility, whereas the number of microfluidization passes markedly improved solubility. The combined effect of heat and pressure significantly increased heat coagulation time. Emulsifying activity index substantially increased upon heat denaturation and was further enhanced by microfluidization. Emulsion stability appeared unaffected by the combined treatment, but the concentration of adsorbed protein on fat droplets was significantly increased. Foaming properties were diminished by heating. Particle size distribution patterns, sodium dodecyl sulfate-PAGE, and size exclusion-HPLC revealed disappearance of major WP and creation of relatively higher, as well as smaller, molecular weight aggregates as a result of the 2 treatments. The use of heat and microfluidization in combination could be used to stabilize WP against heat by producing microparticulated species that have different surface and colloidal properties compared with native WP. These results have implications for the use of WP as an additive in heat-processed foods.

Key words: whey protein, heat denaturation, microfluidization, functional property

 

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Introduction 

Whey proteins (WP) are unique nutritional supplements and functionally valuable food ingredients. They have important biological (digestibility, amino acid pattern, high biological value, and sensory characteristics), physical, and chemical functionalities (McIntosh et al., 1998). Based on their amino acid composition and rate of peptide and amino acid release in the small intestine, WP are nutritionally superior functional food supplements compared with other dietary proteins (Vasiljevic and Shah, 2007). They are extensively used in various food applications, with an increasing global demand. Different types of WP powders are commercially available. Among these, the whey protein concentrates and whey protein isolates are most widely used (Fox and McSweeney, 2003). Whey proteins are highly soluble over a broad range of pH (Zhu and Damodaran, 1994), a property that is important in their application as foaming, emulsifying, gelling, and water-binding agents in various types of food products. Whey proteins are used in sport beverages, liquid meat replacements, baked products, processed meats, salad dressings, ice creams, artificial coffee creams, soups, and various dairy products (Fox and McSweeney, 2003; Fitzsimons et al., 2007.

Whey proteins are composed of a mixture of β-LG, α-LA, BSA, immunoglobulins, proteose peptones, and other minor proteins (Fitzsimons et al., 2007). In their native form they exist as compact, globular proteins (Lee et al., 1992) with high solubility due to a large proportion of surface hydrophilic residues. When subjected to denaturing agents such as heat, previously buried hydrophobic groups become exposed and sulfhydryl/disulfide exchange chain reactions take place between exposed cysteine residues resulting in dissociation, partial unfolding, and finally, aggregation (Lee et al., 1992). The rates and pathways of these physicochemical reactions are determined by environmental factors such as protein concentration, pH, temperature, ionic strength, and solvent condition (Brandenberg et al., 1992; Marangoni et al., 2000; Iordache and Jelen, 2003).

Whey proteins are excellent foaming and emulsifying agents. To obtain optimum foaming and emulsifying characteristics the protein must be substantially soluble, diffuse to the newly formed interface, unfold and reorient in ways that lower interfacial tension, and form cohesive and viscoelastic films by polymerization mainly via disulfide bonds and hydrophobic interactions (Lee et al., 1992; Monahan et al., 1993; Bouaouina et al., 2006). The number of sulfhydryl groups, molecular flexibility, hydrophobicity, and surface activity determine the ability of WP to form stable emulsions and foams (Lee et al., 1992). Although the normal functionality of WP mainly depends on the behavior of β-LG, the most abundant protein in whey (Verheul et al., 1998), overall functionality depends on the combined properties of all WP components. Whereas β-LG has excellent gelling, foaming, and emulsifying properties, α-LA exhibits good emulsifying properties but has poor gelation properties (Pearce and Kinsella, 1978). Molecular flexibility enhances emulsion and foam formation by increasing the rate of unfolding at the interface and allowing more favorable alignment of polar and nonpolar groups in their preferred phase (Klemaszewski and Kinsella, 1991). Sulfhydryl groups, which are more active at pH 7, contribute to molecular flexibility, whereas disulfide bridges play a role in rigidity. It is well known that there is an improved foaming and increased foam stability at pH 7 when WP solutions are first heated to 55°C (Phillips et al., 1990b). At higher temperatures, however, foaming and emulsifying characteristics may be impaired due to protein aggregation, thereby decreasing the availability of protein to form films and stabilize emulsions (Phillips et al., 1990b).

Whey proteins have significant potential use in food manufacturing, but a major obstacle restricting their application is heat-induced destabilization. In industrial practice, the inevitable heat treatment employed when WP are concentrated changes the native state, thus affecting their stability. In addition, compulsory heat treatments during processing of some food products containing WP may cause protein denaturation, aggregation, and flocculation resulting in phase separation, destabilization of emulsions, or protein precipitation (Pearce and Kinsella, 1978; Patel and Kilara, 1990). Thus, it is important for industry to improve the functionality of WP to avoid heat-induced adverse effects.

Attempts to improve the functionality of heterogeneous WP preparations by chemical and enzymatic modifications have met with limited success (Spellman et al., 2005). The use of mechanical force under isothermal or isobaric conditions (i.e., by extrusion or static high-pressure treatment) may be a more useful approach. The application of heat and high shear has been the basis for production of microparticulated products such as Simplesse (NutraSweet Co., Deerfield, IL) and Dairy-Lo (Pfizer Inc., New York, NY) (Akoh, 1998). Similarly, Iordache and Jelen (2003) used heat denaturation and microfluidization to improve the solubility of a WP isolate. The application of heat treatment results in the perturbation of protein size and shape and leads to exposure of hydrophobic sites, aggregation, and consequently, network formation (Gosal and Ross-Murphy, 2000). Under the influence of an applied shear, links between or within the network may break, leading to fragmentation of clusters (Shih et al., 1990). Structural modifications of whey protein preparations by heat treatment and dynamic high-pressure shearing could result in products with distinctly different functional characteristics from those of the native preparations (Huppertz et al., 2005; Considine et al., 2007). The general objective of this study was to assess the effects of dynamic high-pressure shearing (microfluidization) on some functional properties of native and heat-denatured whey protein preparations.

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Materials and Methods 

Materials and Proximate Composition 

Two different batches of WP retentate containing approximately 30% total solids and their corresponding whey permeates obtained on 2 occasions were kindly provided by Warrnambool Cheese and Butter Factory (Warrnambool, Victoria, Australia). Initial compositional analysis of these samples was carried out following established AOAC methodology. Total protein was determined using a Kjeldahl method with a nitrogen conversion factor of 6.38 (AOAC, 2000; method 968.06); moisture was determined by drying (AOAC, 2000; method 925.10); ash content was estimated by combusting presolidified samples (AOAC, 2000; method 923.03); fat content was determined using the Mojonnier method (AOAC, 2000; modified method 989.05); calcium content was determined by atomic absorption spectrophotometry (AOAC, 2000; method 985.33). Phosphorus content was determined colorimetrically using vanadomolybdophosphoric acid (Gales et al., 1966). Lactose concentration was measured by HPLC (Vasiljevic and Jelen, 2003). Results of proximate analysis are given in Table 1.

Table 1. Compositional analysis of whey protein retentates and permeates
ComponentRetentatePermeateSEM1
Moisture, % wt/wt70.1094.780.26
Protein, % wt/wt24.530.860.37
Fat, % wt/wt1.510.050.12
Lactose, % wt/wt1.893.740.04
Ash, % wt/wt1.000.500.03
Ca, mg/L470.21,455.0181.6
P, mg/L309.4658.877.3

1SEM denotes the pooled standard error of the mean, P < 0.05. Means represent the average of at least 6 independent observations (n = 6).

Treatment of Samples 

Upon protein determination, WP retentate samples were adjusted to 10% (wt/wt) protein using the corresponding whey permeates. The resulting solutions had their pH adjusted to 7 with NaOH before being subjected to heat and high-pressure treatments. Two samples were microfluidized (Model 110Y, Microfluidics, Newton, MA) using 1 and 5 passes at 140MPa without heat treatment (N1 and N5); 2 other samples were first heat-denatured at 90°C for 20min for complete WP denaturation (Fox and McSweeney, 2003) and then microfluidized using 1 and 5 passes at 140MPa (H1 and H5). An untreated whey protein concentrate served as the control (C). The samples were then spray dried using a pilot-scale spray dryer (SL-10 Mini-Maxi Pilot Spray Dryer, Saurin Enterprises Pty. Ltd., Melbourne, Australia). All powders were placed in airtight plastic containers and stored at ambient temperature until needed for further analysis.

Particle Size Distribution 

Approximately 1% (wt/wt) WP dispersions were prepared from the spray-dried powders and stored overnight at 4°C to achieve full hydration of proteins. The particle size distributions of control and non-heat-treated samples were determined by a dynamic light scattering instrument (Zetasizer-Nano ZS, Malvern Instruments, Worcestershire, UK) equipped with Dispersion Technology software (version 5, Malvern Instruments). Particle size distributions of heat-treated samples were similarly analyzed by a Mastersizer (Model 2000, Malvern Instruments).

Size Exclusion-HPLC 

The degree of aggregation and denaturation of different WP preparations was observed using size exclusion (SE)-HPLC as described by Bouaouina et al. (2006) with slight changes. The chromatographic separation of WP was performed on an HPLC column (BioSep-SEC-S 3000, 300×7.8mm, Phenomenex, Lane Cove, New South Wales, Australia), and eluted components were detected by UV absorption at 280nm (model 9050, Varian Analytical Instruments, Walnut Creek, CA). Approximately 5% (wt/wt) WP dispersions were diluted in the mobile phase (0.05 M KH2PO4, pH 6.8) to give a protein concentration of approximately 4 mg/mL, and then filtered through a 0.45-μm filter before loading into the column. The solvent flow was maintained at 0.5 mL/min isocratically and the injection volume was 20μL. The resulting chromatograms were analyzed using commercial software (Varian-Star Chromatography Workstation, version 5.31). Standard solutions of α-LA (Sigma-Aldrich Chemie GmbH, Steinheim, Germany), β-LG (Sigma-Aldrich Chemie GmbH), and BSA (Sigma-Aldrich Chemie GmbH) at a concentration of 10 mg/mL were used to quantify WP.

SDS-PAGE 

Protein compositions of each WP preparation were also examined by SDS-PAGE under reducing and nonreducing conditions. Samples were prepared as described by Ong et al. (2006) with some modifications. Approximately 0.01g of WP powder was dissolved in a reducing buffer (1mL of 10mM Tris, 1mM EDTA, pH 8.0, 350μL of 10% SDS, 50μL of β-mercaptoethanol), denatured at 95°C for 5min in a water bath, and diluted 1:4 with buffer (0.125 M Tris-Cl, 4% SDS, 20% vol/vol glycerol, 0.2 M dithiothreitol, 0.02% bromophenol blue, pH 6.8). Protein standards used in the electrophoresis were α-LA and β-LG (Sigma-Aldrich Chemie GmbH; 4mg of each dissolved in 2mL of Milli-Q water, diluted 1:2 in buffer) as well as the broad-range prestained SDS-PAGE standards (161-0318, Bio-Rad Laboratories, Hercules, CA). The WP samples, α-LA, β-LG (6μL), and molecular weight standards (8μL) were then loaded onto 4-20% iGels (NuSep, French Fores, Australia). The gels were placed in a Bio-Rad Protean ll xi cell filled with the tank buffer (0.025 M Tris, 0.192 M glycine, 0.1% SDS, pH 8.3) and run at 50mA for 50min. The gels were then placed in destaining solution 1 (40% methanol, 7% acetic acid) for 30min, stained with staining solution (0.025% Coomassie Brilliant Blue R 250, 40% methanol, 7% acetic acid) for 24h, destained in destaining solution 1 for 1h followed by further destaining in destaining solution 11 (5% methanol, 7% acetic acid) until the background became clear. Gel images were taken using a Fuji Film Intelligent Dark Boxll with Fuji Film LAS 1000 Lite V1.3 software (Fuji Photo Film Co. Ltd., Tokyo, Japan).

Differential Scanning Calorimetry 

Thermal analysis of WP preparations was carried out using a differential scanning calorimeter (DSC 7, Perkin Elmer, Norwalk, CT) equipped with software (Pyris Manager, v.5.0002). The instrument was calibrated for temperature (T) and denaturation enthalpy (ΔH) using indium (Tpeak = 155.87°C, ΔH = 28.234 J/g) and zinc (Tpeak = 417.4°C, ΔH = 93.337 J/g). Approximately 30-μL samples of 12% (wt/wt) WP dispersions were accurately weighed into aluminum pans and hermetically sealed. An empty pan of equal weight was used as a reference. The scanning temperature was raised from 25 to 100°C at 10°C/min. The temperatures at peak height, onset, and endset temperatures and ΔH values were recorded.

Functional Properties of Treated WP 

Analysis of powders for various functional properties was carried out using 5 (% wt/wt) protein dispersions at pH 7 unless otherwise stated. The protein powder needed to make the protein dispersions was weighed into a clean, dry, closed beaker, and Milli-Q water was added with continuous stirring for approximately 30min at room temperature. The dispersion was maintained at 4°C overnight to allow for full hydration of proteins. The pH of the dispersions was adjusted to 7 with NaOH and the final weight was corrected before testing.

Solubility 

Protein solubility was estimated by the method of Morr et al. (1985) with minor modifications. Ten-milliliter portions of protein dispersions were centrifuged (Model J2HS, Beckman, Fullerton, CA) at 12,000 × g at 20°C for 20min, and the supernatant was filtered through a 0.45-μm filter. The protein content of the supernatant and the original dispersion was estimated by using Bradford reagent (Sigma-Aldrich, St. Louis, MO) using a standard curve (r2 = 0.9848) developed with BSA (Sigma-Aldrich Chemie GmbH). Measurements were taken with a spectrophotometer (Novaspec ll, Pharmacia LKB, Norfolk, UK) and percentage solubility was calculated using the following equation:

Heat Stability 

Heat stability of WP was assessed at 140°C by 2 different methods: 1) heat coagulation time (HCT; Rattray and Jelen, 1996) and 2) solubility after brief exposure to heat. Heat coagulation time is defined as the time required to observe formation of visible aggregates during exposure to excessive heating. Precisely measured 3.0-mL samples of WP dispersions were placed in glass tubes (10-mm diameter and 75-mm length), sealed, immersed, and rocked in an oil bath (shaking water bath, Ratek, Boronia, Australia) at 140°C. The time when the first visible aggregates appeared was recorded as HCT. For the solubility method, 3-mL samples of WP dispersions were placed in similar sealed glass tubes and exposed to the same conditions for 10s. Then, tubes were quickly removed from the oil bath, cooled instantly in an ice bath, and centrifuged (Model J2HS, Beckman) at 12,000×g and 20°C for 20min. Samples of original protein dispersions were also centrifuged, the supernatants of both instances were filtered through 0.45-μm filter, and the protein content of the filtered supernatants was measured as before in solubility. Heat stability was expressed using the following equation:

Emulsifying Activity Index and Emulsion Stability Index 

Protein dispersions were analyzed by a turbidimetric technique for emulsifying activity index (EAI), emulsion stability index (ESI), and adsorbed protein as described by Pearce and Kinsella (1978) with some changes. Emulsions of protein dispersions were prepared by mixing 80.0mL of canola oil with 240.0mL of 5% (wt/wt) WP dispersions. This mixture was held in a water bath at 50°C for 20min, homogenized initially by a homogenizer (Polytron, Kinematica AG. Lucerne, Switzerland) for 3min at 50°C and then microfluidized (Model 110Y, Microfluidics) using 1 pass at 140MPa at room temperature.

The EAI was calculated using the following equation expressed as units of area of interface stabilized per unit weight of protein:

where T denotes turbidity; Φ is the oil volume fraction; and C is the weight of protein per unit volume of aqueous phase before an emulsion is formed.

Aliquots (1mL) of the emulsions were diluted serially with 0.1% SDS to give a final dilution of 1/3,000. The absorbance of the diluted emulsions was determined in a 1.5-cm path length cuvette at 500nm in a spectrophotometer (Novaspec ll, Pharmacia LKB). In theory, when none of the light scattered by the turbid sample reaches the photodetector in a sample that does not adsorb light at 500nm, turbidity is given by:where A is the observed absorbance and l (m) is the path length of the cuvette (Pearce and Kinsella, 1978).

Aliquots (1mL) of emulsions and protein dispersions were dried separately in an IsoTemp oven (Fisher Scientific, Suwanee, GA) at 120°C to constant weight. The oil volume fraction (Φ) was calculated by using the following formula:

where A is the mass of empty pan; B is the mass of pan plus emulsion; C is the mass of pan plus DM of emulsion; Do is the density of oil; Ds is the density of protein dispersion; and E is the concentration of protein in dispersion (Pearce and Kinsella, 1978).

Emulsions were held at 4°C for 24h before analysis for ESI, which was calculated by the following formula:

where T is the turbidity value at zero hours; ΔT is change in turbidity; Δt is the time interval (Pearce and Kinsella, 1978).

Ten-milliliter aliquots of each emulsion were centrifuged at 12,000 × g (J2HS, Beckman) at 20°C for 30min and the protein content of the aqueous layer was measured using the Bradford reagent (Sigma-Aldrich) as described above. Adsorbed protein was calculated by the equation:

Foam Formation 

Foaming properties of 5% (wt/wt) protein dispersions were examined as reported previously (Phillips et al., 1990a) with minor modifications. Approximately 200-mL portions of protein dispersions were poured into a bowl (3L) of a double beater Sunbeam Mixmaster Compact Pro-400W twin-motor stand/hand mixer (Sunbeam Co. Ltd., Botany, New South Wales, Australia) and whipped for 20min at ambient temperature. The beater rotational speed was set at its highest speed. A sample of foam was quickly and gently added to a 100-mL scoop with a rubber spatula, taking care to avoid entrapped air pockets. Excess foam was scraped off the top of the scoop using a metal spatula to achieve a constant volume for each measurement and the weight of foam was recorded. This part of the procedure was conducted within 1min. The foam overrun was calculated by the following equation:

Foam stability was measured by monitoring drainage at ambient temperature (Phillips et al., 1990a). To facilitate continuous measurement of drainage from foams, the stainless steel whipping bowl (3L) was modified by drilling a 0.6-cm hole in the bottom, 5.1cm from the center. The edge of the hole was located outside the path taken by the outside beater. The hole was sealed during whipping by adhesive tape applied to the outside of the bowl. The foams were generated from 200mL of protein dispersion. The beaters, bowl, and the protein dispersion were weighed before whipping for precisely 20min. The bowl, beaters, and foam were quickly weighed to quantify moisture loss during whipping and obtain an accurate weight of liquid in the foam. The tape was then quickly removed, the hole cleared with a glass rod (if necessary), and a bowl placed in a ring stand at a 30° angle above a tared container on a balance pan, so that the hole was at the lowest point. The beaters were oriented to avoid any disturbance of liquid flow. The drained liquid was collected in the tared container on the balance pan and the increase in weight was continuously recorded in 20-s intervals. The time to attain 50% drainage was used as an index of foam stability (Phillips et al., 1990a).

Statistical Analysis 

The study was arranged as a randomized block full factorial design with heat treatment and several microfluidizing passes as the major factors and the replications as blocks. All experiments were replicated twice with subsequent subsampling. Results were analyzed using a GLM (SAS Institute, Cary, NC). The level of significance was preset at P = 0.05.

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Results and Discussion 

Effect of Microfluidization and Heat Treatment/Microfluidization on WP 

The extent of WP denaturation and modification due to heat and high pressure shearing was examined by particle size analysis, SE-HPLC, reducing and nonreducing SDS PAGE and differential scanning calorimetry (DSC). The whey protein samples were of 5 main types: native control (C); native, microfluidized with 1 pass (N1); native, microfluidized with 5 passes (N5); heated then microfluidized with 1 pass (H1); and heated then microfluidized with 5 passes (H5).

Sodium-dodecyl-sulfate PAGE electrophoretograms of different WP preparations under reducing and nonreducing conditions are presented in Figure 1, A to C. The broad-range molecular weight markers and α-LA and β-LG standards were used to identify the protein bands. Generally, under reducing conditions in the presence of β-mercaptoethanol, disulfide bonds are cleaved, and consequently aggregated proteins appear in their monomeric forms. Therefore, as shown in Figure 1A and 1B, protein bands corresponding to heat-treated or non-heat-treated samples showed no substantial change in intensity. Under nonreducing conditions, dissociation of noncovalent bonds takes place (Havea et al., 1998; Patel et al., 2005); thus, WP aggregated via disulfide bonds would remain intact. As expected under nonreducing conditions, corresponding main protein bands (monomeric) disappeared in the heat-treated samples (Figure 1C, sample H1). However, using several microfluidizing passes resulted in reappearance of these bands (Figure 1C, H5), implying possible aggregate fragmentation.

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  • Figure 1. 

    Effects of heating and microfluidization on extent of whey protein denaturation observed by reducing (panels A and B) and nonreducing (panel C) Sds-Page. MWM = molecular weight standard; C = control; N1 = native, 1 pass; N5 = native, 5 passes; H1 = denatured, 1 pass; H5 = denatured, 5 passes.

Figure 2 represents the effect of complete heat denaturation and microfluidization at 140MPa on the particle size distribution of WP powders. The average particle size of denatured and microfluidized WP (samples H1 and H5) was around 10 μm compared with that of non-heat-treated samples (C, N1, and N5) with size distribution between 0.01 and 1 μm. Increasing the extent of high-pressure treatment also widened the particle size distribution of non-heat-treated samples (sample N5). As described by Considine et al. (2007), high hydrostatic pressure induces loss of partial molar volume of globular WP. The compressibility of protein depends on the type of protein and the extent of the treatment and may affect its particle size. High pressure is also known to induce protein denaturation by altering the delicate equilibrium between the interactions that stabilize the folded conformation of native proteins (Considine et al., 2007). The application of dynamic high pressure may also have influenced the conformational rearrangement of proteins (Iordache and Jelen, 2003). Therefore, the denaturation of WP may have occurred mainly because of the loss of cavity volumes, which consequently reduced stability of the hydrophobic core. If the globular proteins were unfolded even to a limited extent, then aggregation of molecules via disrupted hydrophobic groups would most probably result in bigger particles as shown by the change in distribution.

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  • Figure 2. 

    Particle size distribution pattern of whey proteins as affected by heat treatment and microfluidization. N1 = native, 1 pass; N5 = native, 5 passes; H1 = denatured, 1 pass; H5 = denatured, 5 passes.

The SE-HPLC chromatograms showed (Figure 3) that the control and non-heat-treated/microfluidized samples had major peaks corresponding to α-LA, β-LG, and BSA. Also, the major WP were affected by the high-pressure treatment, which decreased their concentrations significantly (P<0.05) with increasing numbers of microfluidization passes (results not shown). As described by Bouaouina et al. (2006), these changes could be expected because of possible conformational rearrangements in WP resulting from the high-pressure treatment. Meanwhile, the peaks corresponding to the major WP were absent in the SE-HPLC chromatograms of heat-treated samples. Further, some larger and smaller molecular weight species were observed in heat-denatured samples, but their concentrations were relatively low. In addition, the aggregates formed during treatment could not have passed the guard column to enter the column because of their size.

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  • Figure 3. 

    Size exclusion-HPLC profiles of native and treated whey protein preparations. C = native control; N1 = native, 1 pass; N5 = native, 5 passes; H1 = denatured, 1 pass; H5 = denatured, 5 passes.

The behavior of WP during heating is accompanied by a change in thermal properties. These enthalpy changes associated with protein unfolding are proportional to the extent of denaturation (De Wit, 1990) and can be observed by DSC. In Figure 4, all non-heat-treated WP samples show one broad endothermic peak with peak height related to denaturation temperature of the major WP, β-LG, which was around 77°C. During WP denaturation the endothermic total enthalpy is related mainly to the disruption of internal hydrogen bonds in protein and water and, to a lesser extent, to the formation of protein-water bonds, excess hydrogen bonds in water around apolar groups, and the disruption of van der Waals bonds between apolar groups (Paulsson and Dejmek, 1990). The ΔH values obtained for the C, N1, and N5 samples were 2.53, 1.95, and 2.20 J/g, respectively. Enthalpy values have been correlated with the content of ordered secondary structure and could be used to monitor the proportion of undenatured protein (Patel et al., 1990). Decrease in denaturation enthalpy of microfluidized samples may be an indication of destruction of hydrophobic interactions caused by high pressure. Rupturing of hydrophobic bonds is an exothermic process (Fennema, 1996) that might be lead to a decrease in ΔH of protein denaturation. The relatively slight increase in ΔH with increasing pressure treatment (N5) may be because of the possible reformation of intra- and intermolecular hydrophobic bonds from resulting disordered protein molecules. In contrast, no such endothermic peaks were detected in heat-treated samples, confirming the prior irreversible denaturation of WP. It presumably reflects the heat stability of those samples, which is consistent with the results of extensively longer HCT.

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  • Figure 4. 

    Differential scanning calorimetry thermograms of 12% (wt/wt) whey protein dispersions. C = control; N1 = native, 1 pass; N5 = native, 5 passes; H1 = denatured, 1 pass; H5 = denatured, 5 passes.

Effect of Treatments on Functional Properties, Solubility, and Heat Stability 

Table 2 indicates that the heat treatment significantly (P<0.05) decreased the solubility of WP (H1 and H5) compared with the control. Native WP are globular with higher numbers of surface hydrophilic residues and buried hydrophobic and cysteine groups resulting in high aqueous solubility (Zhu and Damodaran, 1994; Fox and McSweeney, 2003). β-Lactoglobulin, α-LA, and BSA have 5, 8, and 35 cysteine residues, respectively. Naturally, around neutral pH the intramolecular disulfide bonds between cysteine residues stabilize the tertiary structure of these globular proteins. β-Lactoglobulin exists as a dimer with 2 disulfide bridges and a buried free thiol group; α-LA is a monomer with 4 disulfide bridges but no thiol groups; and BSA is a monomer having 17 disulfide bridges and 1 thiol group (Paulsson and Dejmek, 1990; Monahan et al., 1993). As a result of heat treatment (at 90°C for 20min), their globular conformation is irreversibly changed to a more random structure by exposure of previously buried hydrophobic groups, free thiol oxidation, and sulfhydryl/disulfide interchange reactions between exposed cysteine residues leading to protein aggregation and precipitation. During the application of dynamic high pressure, the induced phenomena of cavitation, shear, turbulence, and temperature increase occur simultaneously (Bouaouina et al., 2006). This may cause conformational rearrangements in the quaternary and tertiary structures of proteins (Bouaouina et al., 2006) with consequent changes of some functional properties such as significantly (P<0.05) increased solubility (sample N1 vs. C) and a significant (P<0.05) decrease in solubility of sample N5 compared with sample N1 as observed in this experiment. In addition, as described by Iordache and Jelen (2003), high-pressure shearing can disrupt the microstructural aggregates formed on heating and cause redispersion.

Table 2. Colloidal and interfacial properties of different whey protein preparations produced using heat and high-pressure shearing
Property2
Treatment1Passes, nSolubility, %Heat stability, %HCT, sEAI, m2/gESI, hAdsorbed protein, mg/mLOverrun, %Foam stability, s
Control096.5b100.5b14.5a5,188a23.3abc32.1a275b0a
N1101.0c99.1b18.9b4,654a23.2bc31.8a1,187d23.7a
594.7b100.0b17.4b4,666a22.3c33.4b850c301.8b
H128.0a93.4a87.8c8,545b24.0ab41.2c0a0a
531.2a89.1a102.5d9,257c24.4a41.2c0a0a
SEM3 1.31.91.13189.070.40.143415.0

a–dMeans with different superscripts in a column indicate significant difference (P < 0.05).

1Treatments: N = non-heat-treated and H = heat-treated whey proteins.

2HCT = heat coagulation time; EAI = emulsion activity index; ESI = emulsion stability index.

3SEM = pooled standard error of the mean, P < 0.05 (n = 8).

As shown in Table 2, heat treatment had a significant (P<0.05) negative effect on heat stability of WP. The comparatively low heat stability in heat-treated samples (H1 and H5 vs. C, N1, and N5) may be due to possible further coagulation and precipitation of protein molecules that have already been denatured or denaturation of remaining native proteins with subsequent precipitation at that temperature. However, the combined effect of heat and number of passes significantly (P<0.05) increased the HCT observed at 140°C (Table 2; H1 and H5). As described earlier native whey proteins are readily denatured upon heating. But heat-treated samples may withstand heat because they have already been denatured and mostly contain no active sites such as free thiol groups to initiate denaturation, which in turn retards heat coagulation. On the other hand, high pressure has a disruptive effect on intramolecular hydrophobic and electrostatic interactions, which finally leads to the subsequent reformation of intra- and intermolecular bonds within or between protein molecules (Bouaouina et al., 2006). Therefore, microfluidization may have increased the interactions of protein molecules, further reducing availability of reactive sites, and consequently increasing the heat stability.

EAI and ESI 

The EAI was significantly (P<0.05) increased (Table 2) by heat treatment (H1 and H5 vs. C, N1, and N5). The number of passes further increased it for denatured samples (H5 vs. H1). In addition, the combined effect of heat and number of passes significantly (P<0.05) increased the concentration of adsorbed protein on the surface of oil droplets (Table 2). The emulsifying activity index is a function of oil volume fraction, protein concentration, and the type of equipment used to produce the emulsion (Pearce and Kinsella, 1978). Generally, heat may reduce the emulsifying characteristics of proteins because of irreversible protein denaturation; however, partial protein unfolding would improve its interfacial properties and thus emulsifying ability (Phillips et al., 1990b). Both surface hydrophobicity (which affects the affinity of the protein for the oil-water interphase) and molecular flexibility (which influences the ability to unfold and interact with other proteins) are important in determining emulsifying activity (Monahan et al., 1993). However, our results revealed an increase in the emulsifying activity in heat-denatured samples. Therefore, the affinity between protein and the dispersed phase might be greater than that of protein-protein, thus imparting a thermodynamically more favorable condition to form stable emulsions. This situation would have been assisted by the method used for emulsion preparation during homogenization of emulsions using dynamic high pressure at 140MPa. The capacity of protein to stabilize emulsions is related to the interfacial area that can be coated by proteins (Pearce and Kinsella, 1978). The exposure of buried hydrophobic groups upon heat treatment may have caused this enhanced emulsification. In addition, microfluidization may have further increased the emulsion properties of heated samples by dispersing microaggregates and changing surface properties by unmasking hidden hydrophobic residues and repositioning them toward oil phase. All emulsions produced in this study had similar (P>0.05) emulsion stability regardless of treatments. Emulsion stability depends on the consistency of the interphase, which does not change with time. Emulsions with an appropriate pH and increased net negative charge present a barrier to the close approach of droplets, thus retarding the rate of coalescence and resulting in more-stable emulsions (Klemaszewski and Kinsella, 1991). In our study, the experiments were carried out at neutral pH. At this pH, WP have a net negative charge, which, in turn, imparts greater emulsion stability by retarding coalescence through repulsion. Also, the reactivity of free thiol groups in undenatured WP is greater at pH 7, which favors the unfolding of protein molecules by initiating sulfhydryl-disulfide interchange chain reactions (Klemaszewski and Kinsella, 1991; Monahan et al., 1993).

Foaming Capacity and Foam Stability 

Results also showed (Table 2) that the foaming properties of WP were detrimentally affected by heating (H1 and H5 vs. C, N1, and N5), whereas the number of passes significantly (P<0.05) increased the foam overrun (N1 vs. C) and stability (N5 vs. C and N1). The extensive aggregation of WP caused by heat denaturation may have reduced the ability of proteins to produce stable films. However, none of the non-heat-treated WP samples (N1 and N5) along with controls showed good foaming ability. Lipids in WPC can seriously impair the foaming ability because surface-active polar lipids interfere with protein films by situating themselves at the air/water interface (Fennema, 1996). Proximate analysis (Table 1) revealed considerable fat and phosphate content in our samples. In addition, these interfering substances possess weak cohesive and viscoelastic properties to overcome the internal pressure of air bubbles compared with WP. As a result, bubbles expand and finally collapse rapidly resulting in poor foaming. According to our results, dynamic high-pressure shearing positively affected both foam overrun and stability. The foaming properties of WP concentrates are significantly correlated with the amount of β-LG (approximately 50% of the total WP; Fitzsimons et al., 2007) and the extent of WP denaturation (Phillips et al., 1990b). As shown in previous studies, β-LG was the most pressure-sensitive WP, and static high pressure of 100 to 200MPa could initiate its structural changes (Considine et al., 2007). Therefore, native microfluidized WP preparations might show improved foaming because of their enhanced molecular flexibility and increased surface hydrophobicity after high-pressure shearing.

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Conclusions 

Combined heat treatment and high-pressure shearing produced micro-aggregates of WP with enhanced heat stability. However, the particle size of these aggregates was large enough to accelerate sedimentation, which in turn resulted in poor solubility. The amount of adsorbed proteins on the surface of fat droplets also increased with combined treatment, which consequently increased emulsifying activity index. High-pressure shearing alone improved foaming properties of native WP. In addition, high-pressure shearing fragmented denatured WP into smaller entities. Microfluidization could be a useful method for stabilizing WP against heat-induced changes by producing microparticulated species that have different surface and colloidal properties.

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Acknowledgments 

Authors gratefully acknowledge the financial and technical support from Victoria University. The Warrnambool Cheese and Butter Factory (Warrnambool, Victoria, Australia) is sincerely appreciated for the provision of required raw materials. We also thank Dairy Innovation Australia Ltd. for partial financial support, Tim Wooster from Food Science Australia (Melbourne, Australia) for technical assistance, and Gwyn Jones for valuable comments during manuscript preparation.

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PII: S0022-0302(09)70449-7

doi:10.3168/jds.2008-1791

Journal of Dairy Science
Volume 92, Issue 4 , Pages 1387-1397, April 2009

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