Food Hydrocolloids 25 (2011) 887e897
Contents lists available at ScienceDirect
Food Hydrocolloids journal homepage: www.elsevier.com/locate/foodhyd
Modiﬁcations of soy protein isolates using combined extrusion pre-treatment and controlled enzymatic hydrolysis for improved emulsifying properties Lin Chen a, Jianshe Chen b, Jiaoyan Ren a, Mouming Zhao a, * a b
College of Light Industry and Food Science, South China University of Technology, 381 Wushan Road, Guangzhou 510641, China School of Food Science and Nutrition, University of Leeds, Leeds LS2 9JT, UK
a r t i c l e i n f o
a b s t r a c t
Article history: Received 3 May 2010 Accepted 18 August 2010
Effects of combined extrusion pre-treatment and controlled enzymatic hydrolysis on the physicochemical properties and emulsifying properties of soy protein isolates (SPI) have been investigated. Results showed that extrusion pre-treatment caused a marked improvement in the accessibility of SPI to enzymatic hydrolysis, resulting in changes in degree of hydrolysis (DH), protein solubility (PS), surface hydrophobicity (H0) and molecular weight distributions (MWD) for ESPIH (extrusion pre-treated SPI hydrolysates). It was observed that emulsion systems formed by control SPI or SPIH (SPI hydrolysates) (20% v/v oil, 1.6% w/v emulsiﬁer, and pH 7.0) were unstable over a quiescent storage period of 21 days, due to bridging ﬂocculation and creaming. However, ESPIH (9.1% DH) was capable of producing a very ﬁne emulsion (d32 ¼ 0.42 mm, d43 ¼ 2.01 mm) which remained stable over a long term quiescent storage. Various surface properties of ESPIH products have also been studied in relation to DH and emulsifying functionalities. It was suggested that signiﬁcantly increased protein solubility and decreased molecular weight could be the main reasons for the greatly improved emulsifying capability of ESPIH. This study demonstrated that modiﬁed soy protein could be an excellent emulsifying agent for food and other applications. It also demonstrated that combined extrusion pre-treatment and enzymatic hydrolysis could be a highly effective method for functionality modiﬁcation of globular proteins. Ó 2010 Elsevier Ltd. All rights reserved.
Keywords: Soy protein isolates Enzymatic hydrolysis Extrusion pre-treatment Protein modiﬁcation Emulsifying properties
1. Introduction The use of soy proteins has been of increased interest, primarily attributed to its high nutritional value, steady supply, and low cost compared to other sourced proteins. Soy protein isolates (SPI), soy protein concentrates (SPC), and soy protein ﬂour (SPF) are the three kinds of widely used soy protein products. Of the three, SPI is often favoured because of its high protein content (ca. 90%) (Nielsen, 1985a). Native soy proteins are composed of a mixture of albumins and globulins, 90% of which are storage proteins with globular structuredconsisting mainly of 7S (b-conglycinin) and 11S (glycinin) globulins (Nielsen, 1985b). The most important functional uses of SPI are as an emulsifying or gelling agent (Kinsella, 1979; Liu, Lee, & Damodaran, 1999; Molina, Papadopoulou, & Ledward, 2001). However, because of the compact globular structure stabilized mainly by hydrogen bonds and disulﬁde bonds, native soy proteins have lower molecular ﬂexibility and a rather poor emulsifying capability compared to other protein emulsiﬁers such as milk
* Corresponding author. Tel.: þ86 208 7113 914; fax: þ86 208 7114954. E-mail address: [email protected]
(M. Zhao). 0268-005X/$ e see front matter Ó 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.foodhyd.2010.08.013
protein (Molina et al., 2001; Roesch & Corredig, 2003; Staswick, Hermodson, & Nielsen, 1984). In particular, commercial soy protein products undergo harsh processing conditions including spray drying and high temperature sterilization, and may lose much of their functionalities (Lee, Ryu, & Rhee, 2003; Wagner, Sorgentini, & Añón, 2000). Emulsifying capability and emulsion stability are considered to be two different attributes that are inﬂuenced by different physicochemical properties of the proteins or peptides (Walstra & Smulders, 1997). During the formation of an oil-in-water emulsion in a homogenizer, oil droplets are broken up into smaller ones by intense laminar ﬂow (shear and extensional deformations) and/ or the inertial effects (turbulence and cavitations) and surfaceactive agents adsorb to the newly created surfaces to protect droplets against bridging ﬂocculation or re-coalescence. Proteins or peptides aid in the formation of oil droplets, mainly by lowering interfacial tension (facilitating droplet breakup) and by forming a physical barrier at the oilewater interface to prevent the bridging ﬂocculation and re-coalescence of droplets (Walstra, 1993). The ﬁnal size of droplets produced during homogenization depends on two processes: (i) the initial generation of droplets of small size, and (ii) the rapid stabilization of these droplets against bridging
L. Chen et al. / Food Hydrocolloids 25 (2011) 887e897
ﬂocculation and re-coalescence once they are formed (McClements, 2005). Emulsifying capability therefore refers to the ability of an emulsiﬁer to form and stabilize small droplets during homogenization. Once emulsions are formed, they are subjected to several forms of instability, including Ostwald ripening, creaming, ﬂocculation, and coalescence. The latter three forms of instability were the main concern in this work. Emulsion stability is favoured by proteins and peptides because of their abilities in opposing attraction between emulsion droplets by promoting electrostatic repulsion or steric hindrance (Damodaran, 2004). Because of low side reactions and products, enzymatic hydrolysis has been widely used to improved protein functionalities (Govindaraju & Srinivas, 2006; Tsumura, 2009). Physico-chemical properties of protein hydrolysates mainly depend on the degrees of hydrolysis (DH) and types of protease used (Adler-Nissen, 1986). Many studies have demonstrated that controlled enzymatic hydrolysis of globular protein can improve their emulsifying properties, by improving overall protein solubility, exposing hidden hydrophobic residues, and reducing the molecular size (Radha & Prakash, 2009; Tsumura, 2009). Controlled enzymatic hydrolysis means that the hydrolysis modiﬁcation was regulated with a combination of temperature, time, enzyme concentration, etc for a desirable DH value. However, some studies showed that excessive enzymatic hydrolysis could be detrimental to their emulsifying properties (Qi, Hettiarachchy, & Kalapathy, 1997; Wu, Hettiarachcchy, & Qi, 1998). Pancreatin, mainly containing trypsin and chymotrypsin, is produced by the exocrine cells of the pancreas and has its maximum activity at alkaline pH (Qi et al., 1997). Because of the very broad speciﬁcity to peptide bonds, pancreatin has often been used to investigate the digestibility of proteins (Kane & Miller, 1984; Rothenbuhler & Kinsella, 1986). However, previous studies have found that soy proteins appeared to be less accessible to enzymatic hydrolysis (Marsman, Gruppen, Mul, & Voragen 1997; Tsumura, Saito, Kugimiya, & Inouye, 2004). Because of the compact quaternary and tertiary structures in soy protein, many hydrolysis sites are shielded and difﬁcult to be accessible by protease. Recently, several studies reported that extrusion pre-treatment could improve the accessibility of globular protein to _ ski, Fik, enzymatic hydrolysis. For example, Surówka, Zmudzi n Macura, and qasocha (2004) demonstrated that when the same enzyme: substrate (E/S) ratios (ratios of enzyme to substrate concentrations) were introduced, extruded SPF attained higher DH values than untreated SPF did. Furthermore, they found that soy glycinin, the subunit of soy protein that was resistant to enzymatic hydrolysis, can be completely hydrolyzed using Protamex and Neutrase after extrusion pre-treatment. Marsman et al. (1997) also found that after extrusion pre-treatment, all the subunits in soy proteins can be rapidly and completely degraded with Neutrase. It has been suggested that during extrusion, the combined effect of high temperature, high pressure and shear forces in the extruder leads soy proteins to form laminated structures, which were easily accessible to _ ski, 2004). enzymes (Surówka & Zmudzi n Combined extrusion pre-treatment and enzymatic hydrolysis can cause changes in the physico-chemical properties and functionalities of soy protein hydrolysates. However, ambiguous or sometimes contradictory results were reported in literature. Surówka et al. (2004) demonstrated that the combined extrusion pre-treatment and controlled enzymatic hydrolysis using Neutrase could increase the emulsifying activity index (EAI) of SPC, while Jung, Murphy, and Johnson (2006) observed that such a modiﬁcation caused a decrease in EAI and an increase in emulsion stability index (ESI) for SPF. One of the main limitations of these studies, however, was that emulsifying properties were mainly characterized by EAI and ESI using turbidity measurement, but not the fundamental mechanisms underpinning the emulsion formation
and stabilization. Therefore, a systematic study is needed to identify the inﬂuence of the changes in physico-chemical properties and interfacial properties of soy protein hydrolysates caused by extrusion pre-treatment on their emulsifying properties. But to our knowledge, no such study has been reported so far in literature. The aim of the present work was to study the effects of combined extrusion pre-treatment and controlled enzymatic hydrolysis using pancreatin on the emulsifying properties of SPI. It was also hoped that by measuring interfacial properties, the underpinning mechanisms of improved emulsifying properties for SPIH and ESPIH can be clariﬁed. 2. Materials and methods 2.1. Materials and chemicals SPI was obtained from Wonderful Industrial Group (Shandong Province, China). The crude protein content of SPI determined by micro-Kjeldahl method (N 6.25) was 93.24% 0.73 (w/w). The moisture content was 5.13% 0.18 (w/w). Pancreatin powder (from porcine pancreas) was purchased from Hangzhou Sanye Chemical Co. (Hangzhou, China). This enzyme product is a mixture of trypsin (EC 3. 4. 21. 4), chymotrypsin (EC 3. 4. 21. 1), elastase (EC 3. 4. 4. 7) and carboxypeptidase A (EC 3. 4. 17. 1). According to manufacturer, it had a speciﬁc activity of 3000 BAEE (N-benzoyl-L-arginine ethyl ester) units/mg, where BAEE unit is the standard unit for the activity of protease. One BAEE unit will produce a DA253nm of 0.001 per min at pH 7.6 at 25 C using BAEE as substrate. 1-Anilino-8naphthalenesulfonate (ANS) reagent was purchased from Sigma (St. Louis, MO, USA). All other chemicals used in the present study were of analytical grade. Millipore water (water puriﬁed by treatment with a Milli-Q apparatus, Millipore, Bedford, UK) was used for the preparation of the solutions. 2.2. Extrusion pre-treatment of SPI SPI was subjected to extrusion in a laboratory-scale twin-screw extruder (SPJ-40, Deai Co., Ltd, Shanxi, China) with three individual barrel sections, each with separate temperature control. The temperature of these three barrels was set at 50 C (feed section), 90 C, and 160 C, respectively. The diameter of the screw was 75 mm, and the length-to-diameter ratio was 28:1. The screw elements included kneading blocks and reverse screw elements. Screw speed was operated at 220 rpm. The die was designed with two circular holes at 5 mm diameter. The moisture content of SPI was adjusted to 20% speciﬁed for the extruder. Moisturised raw material was introduced to the extruder at a rate of 0.2 kg/min. The obtained extrudates of SPI (ESPI) were ground to pass a 0.2 mm screen and then oven-dried at 50 C to reach a ﬁnal moisture content of 5.1% (w/w). Ground ESPI was stored in air-tight glass containers. 2.3. Preparations of SPIH and ESPIH with various DH values Hydrolysates with various DH values were prepared from 1.6% (w/v) SPI and ESPI dispersions by hydrolysis at pH 7.0 and 55 C using pancreatin, where the concentration was calculated based on the amount of powder per 100 mL solution (same calculation was used throughout this work). Hydrolysates were freshly prepared prior to each experiment. The pH and DH were controlled using the pH-stat method by using a TIM840 Auto titrator (Radiometer Analytical Co., USA) (Adler-Nissen, 1986). The E/S ratios (g enzyme/ g substrate) varied from 0.01 to 5.0% (w/w), and the molarity of the NaOH solution used to maintain the pH varied from 0.1 to 2.0 M. In preliminary experiments, DH as a function of the E/S ratio and
L. Chen et al. / Food Hydrocolloids 25 (2011) 887e897
hydrolysis time was investigated for both SPIH and ESPIH. Based on the investigations of preliminary experiments, the appropriate E/S ratios were selected to reach different required DH values for both SPIH and ESPIH. And the hydrolysis time was determined to be 180 min, wherein the progression of hydrolysis can reach a plateau. After hydrolysis, the enzymes were inactivated by placing in boiling water for 15 min. Heating of the hydrolysate dispersions resulted in a slight decrease in pH. 0.1 M NaOH was used to readjust the hydrolysate dispersions to pH 7.0. The control SPI and control ESPI dispersions were prepared under the same incubation conditions and heat inactivation treatment, but without pancreatin adding. Samples codes are subsequently composed of the name of protein source and the DH reached. For example, SPIH 1.0% means soy protein isolates hydrolysate with DH 1.0%. DH is deﬁned as the ratio of the number of peptide bonds cleaved (number of free amino groups formed during proteolysis) expressed as hydrolysis equivalents (h), in relative to the total number of peptide bonds before hydrolysis (htot).
DH ð%Þ ¼
h 100% htot
In this work, the DH for enzymatic hydrolysis was measured by the pH-stat method. The percent DH was calculated according to the following equation:
DH ð%Þ ¼
a Mp htot
where B is the base consumption in mL, Nb is the normality of the base, a is the degree of dissociation of a-NH2 groups, Mp is the mass of protein being hydrolyzed (g), and htot is the total number of peptide bonds in the protein substrate (meqv/g protein). According to Adler-Nissen (1986), htot was 7.75 (meqv/g protein) and a was 0.44 for SPI. 2.4. Characterization of physico-chemical properties of SPIH and ESPIH 2.4.1. Protein solubility (PS) measurement Protein solubility (PS) was determined according to the method of Petruccelli and Añón (1994), with minor modiﬁcations. 100 mL of SPIH and ESPIH dispersions (1.6%, w/v) were centrifuged at 12,000 g for 30 min at 20 C in a CR22G centrifuge (Hitachi Co., Japan). The supernatants were collected. Protein content of the supernatants was determined by micro-Kjeldahl method (N 6.25). Protein solubility was calculated as nitrogen solubility index (NSI, %) ¼ (protein content of supernatant/amount of proteins added) 100%. 2.4.2. Surface hydrophobicity (H0) measurement Surface hydrophobicity (H0) was determined by the hydrophobicity ﬂuorescence probe 1-anilino-8-naphthalenesulfonate (ANS) according to the method of Kato and Nakai (1980), with minor modiﬁcations. Dispersions of selected SPIH and ESPIH were freezedried and dispersed into buffer solution (0.01 M phosphate buffer, pH 7.0) to reach different concentrations (0.004e0.02%, w/v), followed by stirring at 4 C overnight. After centrifugation (12,000 g, 20 min, 20 C), the supernatants were collected. Then, aliquots (20 mL) of ANS (8 mM in the same buffer) were added to 4 mL of sample. Fluorescence intensity (FI) was measured at wavelengths 390 nm (excitation) and 470 nm (emission) using a RF-5301 PC spectro-ﬂuorometer (Shimadzu Corp., Kyoto, Japan) at 26 C, with a constant excitation and emission slit of 5 nm. The initial slope of ﬂuorescence intensity versus protein concentration (mg/mL) was calculated by linear regression analysis and used as an index of H0.
2.4.3. Determination of molecular weight distribution (MWD) The molecular weight distributions of proteins or peptides were determined by size exclusion chromatography (SEC). A protein puriﬁcation chromatography (Amersham plc., Buckinghamshire, UK) with a Superdex75_peptide_10/300_GL column was used for analysis. The mobile phase was 0.01 M sodium phosphate buffer containing 0.1 M NaCl (pH 7.2) and was degassed and ﬁltered through 0.2 mm ﬁlter before used. Dispersions of selected SPIH and ESPIH with various DH values were freeze-dried and dispersed into the mobile phase to a ﬁnal concentration of 3 mg/mL, followed by stirring at 4 C overnight. After centrifugation (12,000 g, 20 min, 20 C), the supernatants were ﬁltered on a 0.2 mm membrane (PVDF). 100 mL of the supernatants was applied to the column. Elution was performed isocratically at a ﬂow rate of 0.5 mL/min. Detection was performed at 220 nm. The column was calibrated with a low molecular weight (MW) calibration kit (product code 28-4038-41, Amersham plc., Buckinghamshire, UK). The MW of the individual proteins included in the 28-4038-41 calibration kit and their respective elution volumes were: Aprotinin (6.5 kDa, 15.64 mL), Ribonuclease A (13.7 kDa, 13.13 mL), Carbonic Anhydrase (29.0 kDa, 11.37 mL), Ovalbumin (43.0 Da, 10.19 mL), Conalbumin (75.0 kDa, 9.69 mL). The following calibration curve was then obtained: Kav ¼ 0.368l g MW þ 1.871 (where Kav is gel-phase distribution coefﬁcient). UNICORN 5.0 software (Amersham plc., Buckinghamshire, UK) was used to collect and analyze the chromatographic data. 2.5. Characterization of emulsifying properties of SPIH and ESPIH 2.5.1. Preparations of emulsions SPI dispersions were prepared by dispersing the desired amount (0.2e10.0%, w/v) of SPI into Millipore water and then gently stirring overnight at ambient temperature. The pH of the resulting protein solution was adjusted to pH 7.0 by adding a few drops of 1 M NaOH or 1 M HCl. Subsequently quoted pH values and emulsiﬁer concentrations referred to those of the aqueous phase before emulsiﬁcation. Sodium azide (0.02%, w/v) was added to the buffer as an anti-microbiological agent. Oil-in-water emulsions containing 20% (v/v) sunﬂower seed oil and 80% (v/v) protein dispersion were prepared at ambient temperature using a single-pass laboratoryscale jet homogenizer operating at the pressure of approximately 300 bar (Burgaud, Dickinson, & Nelson, 1990). 2.5.2. Determination of droplet size distribution and mean droplet size Droplet size distributions (individual droplets or droplet aggregates) of emulsion samples were determined using a Malvern Mastersizer MS2000 (Malvern Instruments Ltd., Worcestershire, UK), by applying the following optical parameters: sunﬂower seed oil and water refractive indices: 1.462 and 1.330, respectively; absorption index: 0.001. The mean droplet size was characterized in terms of the surface area mean diameter d32 and volume mean diameter d43. The d32 value was used to estimate the speciﬁc surface area of freshly made emulsions, and the d43 value was used to monitor changes in droplet size. 2.5.3. Confocal laser scanning microscopy (CLSM) observation A Leica TCS SP2 confocal laser scanning microscope (Leica, Heidelberg, Germany) mounted on a Leica DM-RXE upright microscope base was used to visualize the microstructure of the emulsion samples. A 63 water-immersion objective with numerical aperture 1.20 was used in all of the experiments reported here. The CLSM was operated in ﬂuorescence mode. Emulsion oil phase was stained with Nile Red dye (20 mL of 0.01% (w/v) dye in polyethylene glycol added to 5 mL emulsion). The stained
L. Chen et al. / Food Hydrocolloids 25 (2011) 887e897
emulsions were immediately loaded into a laboratory-made welled slide to ﬁll it completely. The well was 8 mm in diameter and 1.6 mm in depth (volume w 80 ml). Fluorescence from the sample was excited with the 488 nm line of an Argon laser as the light source; the emission peak of the light was around 565 nm. As Nile Red stains the oil phase, individual large oil droplets and regions rich in emulsion droplets appear as bright patches, whereas the aqueous (water/protein) phase appears dark in the microimages. Images were recorded at 24 C at a resolution of 1024 1024 pixels and processed using Leica Qwin software (Leica, Heidelberg, Germany). Eight scans were averaged during the creation of each image. 2.5.4. Emulsion storage stability measurement Emulsion samples were poured into 5 mL glass tubes (height 75 mm, diameter 9 mm) immediately after preparation. Subsequently, the tubes were sealed to prevent evaporation. The emulsion samples were stored quiescently at ambient temperature for 21 days. During storage, the extent of creaming was characterized by creaming index (CI, %): CI ¼ (HS/HE) 100%, where HS is the height of serum layer, and HE is the total height of the emulsion. The changes in mean droplet size were monitored by measuring d43; the changes in microstructure were assessed by examining emulsions using CLSM.
micro-Kjeldahl method (N 6.25). The surface load (Gsat, mg/m2) was calculated as Equation (3):
Ci Ceq d32 ¼ 6Ø
where Ci is the initial protein concentration per unit volume of emulsion (kg/m3) and Ceq is the concentration of the non-adsorbed protein per unit volume of emulsion (kg/m3). In the current study, d32 of emulsion was determined by light scattering (Mastersizer); Ø is oil volume fraction (¼0.2). The fraction of protein adsorbed (Fads, %) onto the droplets during homogenization was calculated as Equation (4):
Ci Ceq 100% Ci
where the Ci and Ceq are the same as that described in Equation (3). 2.7. Statistical analysis Unless otherwise stated, all the tests were conducted in triplicate and the results were expressed as the mean standard deviation. Analysis of variance was conducted to identify differences among means by one-way ANOVA using SPSS 13.0 software (SPSS Inc, Chicago, IL, USA).
2.6. Characterization of interfacial properties of SPIH and ESPIH
2.6.1. Interfacial tension measurement OCA20 video-based contact angle meter (DataPhysics, Berlin, Germany) was used for interfacial tension measurement based on the axisymmetric pendant drop shape analysis. Dispersions of selected SPIH and ESPIH were freeze-dried and dispersed into buffer solution (0.01 M phosphate buffer, pH 7.0) to reach different concentrations (0.001e5.0%, w/v), followed by stirring at 4 C overnight. After centrifugation (12,000 g, 20 min, 20 C), the supernatants were ﬁltered on a 0.2 mm membrane (PVDF) to eliminate the insoluble material. The densities of supernatants and sunﬂower seed oil were detected by using a DMA 35 N density meter (AntonParr, Austria). An aqueous drop of 10 mL was created with the supernatants of SPIH or ESPIH dispersions in a cuvette containing sunﬂower seed oil. Instrument recording started immediately after drop formation. The interfacial tension was studied at ambient temperature for up to 180 min and the data was calculated using the SCA 20 software (DataPhysics, Berlin, Germany). The interfacial pressure (p) was calculated from p ¼ g0 g, where g0 is the interfacial tension at sunﬂower seed oilpure buffer solution interface (g0 ¼ 30.3 0.5 mN/m), and g is the interfacial tension in the presence of the emulsiﬁer. The saturation interfacial pressure (psat) was determined by averaging the measured interfacial pressure values over the range of emulsiﬁer concentrations where p remained relatively constant as the emulsiﬁer concentration was increased further.
3.1. Enzymatic hydrolysis of SPI and ESPI
2.6.2. Determination of protein adsorption fraction (Fads) and surface load (Gsat) The amount of non-adsorbed protein in the continuous phase of the emulsions was determined according to the method described by Akhtar, Dickinson, Mazoyer, and Langendorff (2002), with minor modiﬁcations. Freshly prepared emulsion samples were centrifuged at 12,000 rpm for 2 h at 20 C. After centrifugation, two phases were observed: the cream oil droplets at the top, whereas the serum layer and precipitates at the bottom. The cream phase was carefully removed using a syringe. The amount of non-adsorbed protein remaining in the serum layer and precipitates was determined by
Protein hydrolysis in SPI and ESPI using pancreatin with different E/S ratios was monitored by following DH over 180 min. As shown in Fig. 1A and B, for both SPI and ESPI, enzymatic hydrolysis reaction progressed rapidly at initial stage and then relatively slowly over time before reaching a plateau. This trend is typical for protease hydrolysis. Some physico-chemical properties of SPIH and ESPIH with different DH values used for emulsion experiments are shown in Table 1. It should be noted that when the same E/S ratios were introduced, ESPIH obtained higher DH values than SPIH did. This ﬁnding suggests that pancreatin, which has broad speciﬁcity to peptide bonds, access cleavage sites more easily for ESPI than for SPI. In other words, extrusion pre-treatment used in the current study improved the accessibility of SPI to enzymatic hydrolysis. This ﬁnding agrees with those of several previous studies (Marsman et al., 1997; Surówka et al., 2004). 3.2. Physico-chemical properties of SPIH and ESPIH 3.2.1. Inﬂuence of DH on proteins solubility of SPIH and ESPIH Solubility is one of the most important properties for protein, since it has direct effects on other functionalities. Poor functionality tends to go hand in hand with poor solubility. Because emulsions were prepared at pH 7.0 in the current study, PS of SPIH and ESPIH with various DH values were investigated at this same pH condition. PS of SPIH and ESPIH as a function of DH value at pH 7.0 is shown in Fig. 2. Control SPI showed a poor PS of 15.8%. Because of the harsh processing conditions including spray dry and high temperature sterilization, aggregation of protein may occur in commercial SPI used in the current study, which is detrimental to its PS. After extrusion pre-treatment, PS of ESPI increased to 32.5%. This result suggests that extrusion treatment showed a favourable effect on improving the PS for soy protein, as also reported in the literature (Jung et al., 2006; Surówka et al., 2004). With the increasing of DH value, PS of both SPIH and ESPIH increase
L. Chen et al. / Food Hydrocolloids 25 (2011) 887e897
8 6 4
Fig. 2. Inﬂuence of DH on protein solubility (PS) of SPIH and ESPIH at pH 7.0. Symbols: -, SPIH; >, ESPIH.
controlled enzymatic hydrolysis. This implies that after extrusion pre-treatment, the enzymatic hydrolysis of SPI became not only intensive, but also extensive. This agrees with Surówka and _ ski (2004) ﬁnding that enzymatic hydrolysis of extruded Zmudzi n SPC was more extensive than that of untreated SPC, despite reaching the similar DH. However, it was observed that PS of both SPIH and ESPIH decreased slightly at high DH values. This phenomenon may arise from protein aggregation caused by the increased hydrophobic interaction, eventually leading to the decrease of PS (Wu et al., 1998).
12 9 6 3 0 0
Time (min) Fig. 1. Progression of hydrolysis with time for SPI (A) and ESPI (B) when acted by pancreatin with various E/S ratios. Symbols: C, E/S ¼ 0.1% (w/w); :, E/S ¼ 0.5% (w/w); A, E/S ¼ 2.0% (w/w).
markedly initially, and then decreased slightly at high DH values. It is well known that controlled enzymatic hydrolysis can improve the PS of protein by decreasing molecular weight and increasing charged groups (Panyam & Kilara, 1996). However, we should note that ESPIH showed a larger maximum PS of more than 90% at DH 8.0e10.0%. In contrast, SPIH showed a smaller maximum PS of 69.1% at DH 9.6%. These results demonstrated much improved protein solubility using combined extrusion pre-treatment and
3.2.2. Inﬂuence of DH on surface hydrophobicity of SPIH and ESPIH Protein surface hydrophobicity is an index of the number of hydrophobic groups on the surface of a protein in contact with the polar aqueous environment and is closely related to its emulsifying properties. At a constant pH (e.g. pH 7.0), ANS (a hydrophobicity ﬂuorescence probe) binds to the exposed hydrophobic groups on the surface of a protein, resulting in an increase in ﬂuorescence emission intensity compared with that of free ANS in aqueous solution. And this method was often used to monitor the changes of surface hydrophobicity in protein caused by enzymatic hydrolysis. H0 of SPIH and ESPIH as a function of DH value at pH 7.0 is shown in Fig. 3. Control SPI showed a higher H0 (1109.8) than that of control
Table 1 Properties of SPIH and ESPIH with various DH values used for emulsion experiments. E/S ratio (%, w/w) 0 0.025 0.05 0.1 0.2 0.3 0.5 1.0 2.0
0 0.5 1.0 1.9 2.4 3.0 4.5 6.8 9.6
0 3.1 5.3 8.0 9.1 10.0 11.3 13.3 16.4
15.8 20.2 24.3 30.1 34.4 38.7 43.3 54.3 69.1
32.5 60.6 85.3 90.3a 90.8a 91.6a 85.9 84.0 81.0
1109.8 1437.9 1886.6 736.3 628.4 505.0 194.9 83.8 78.2
323.4 412.3 468.9 457.9b 452.5b 413.6 364.5 253.8 126.7
Results are means of three determinations. Means within a column with the same superscripts are not signiﬁcantly different (p < 0.05).
DH (%) Fig. 3. Inﬂuence of DH on surface hydrophobicity (H0) of SPIH and ESPIH at pH 7.0. Symbols: ,, SPIH; A, ESPIH.
L. Chen et al. / Food Hydrocolloids 25 (2011) 887e897
ESPI (323.4). This indicates that extrusion pre-treatment caused a decrease in H0 for SPI, as also reported by Jung et al. (2006). With the increasing of DH value, H0 of both SPIH and ESPIH increase markedly initially, and then decreased drastically at high DH values. The observed increase in H0 for both SPIH and ESPIH can be explained by the fact that controlled enzymatic hydrolysis of soy protein was accompanied by the exposure of hydrophobic groups in the inner part of the protein. SPIH showed a larger maximum H0 of 1886.6 at DH 1.0%. In contrast, ESPIH showed a smaller maximum H0 of ca. 460 at DH 5.3e9.1%. However, further hydrolysis caused a drastically decrease in H0 for both SPIH and ESPIH. This result may be due to two factors: (1) enzymatic cleavage of hydrophobic bonds that may reside at the surface; (2) protein aggregation caused by the increased hydrophobic interaction (Celus, Brijs, & Delcour, 2007).
Intergrated Area (mAU)
zone I zone II zone III total area
600 400 200 0 Control DH 1.0% DH 1.9% DH 3.0% DH 4.5%
Integrated Area (mAU)
3.2.3. Molecular weight distributions of SPIH and ESPIH Size exclusion chromatography (SEC) was used to study the proteins or peptides MWD of SPIH and ESPIH. As shown in Fig. 4A and B, we found that SEC proﬁles of SPIH and ESPIH can be divided into three zones according to the distributions of chromatogram peaks. They are zone I: <9.69 mL, zone II: 9.69e20.27 mL, and zone Z: 20.27e24.00 mL, corresponding to MWD of >75.0 kDa, 75.0e1.0 kDa, and <1.0 kDa, respectively. The integrated areas of chromatogram peaks for the total or individual zones are shown in Fig. 5A and B, respectively. From Figs. 4 and 5, one can see that most of soluble proteins in control SPI and control ESPI are distributed in zone II (75.0e1.0 kDa).
zone I zone II zone III total area
1500 1000 500 0 Control
DH 3.1% DH 5.3% DH 9.1% DH 13.3%
Fig. 5. Total or individual zone integrated area(s) of chromatogram peaks for SEC proﬁles of SPIH (A) and ESPIH (B) with different DH values. The Roman numerals indicated different zones: zone I, >75.0 kDa; zone II, 75.0e1.0 kDa; zone III, <1.0 kDa.
Fig. 4. SEC proﬁles of SPIH (A) and ESPIH (B) with various DH values. Samples were carried out on a Superdex75_peptide_10/300_GL column, eluted with 0.01 M phosphate buffer (pH 7.2) containing 0.1 M NaCl. Elution volumes of protein standards with MW 75.0 kDa, 43.0 kDa, 29.0 kDa, 13.7 kDa, and 6.5 kDa are indicated from left to right with þ symbols. The Roman numerals indicated different zones: zone I, >75.0 kDa; zone II, 75.0e1.0 kDa; zone III, <1.0 kDa.
Upon hydrolysis, the chromatogram peaks in zone II for both SPIH and ESPIH showed a tendency of shifting towards higher elution volumes (smaller molecular weight). This suggests that enzymatic hydrolysis will cause a decrease in MWD for soy protein, as also reported in literature (Achouri & Zhang, 2001). In SEC proﬁles of all SPIH and ESPIH, there was single one peak eluted at around 20.8 mL in zone Z, corresponding to the MW of 0.55 kDa. This fraction was probably composed mainly of free amino acids. We can see that the area of this peak increased with the increasing of DH value. After eluting with one column volume of mobile phase buffer, all of the solute should be washed away from the column. But in the chromatograms of both SPIH and ESPIH, a new peak at around 31.1 mL was detected. This unusual size exclusion behavior may be due to the hydrophobic interactions of some low molecular weight peptides in hydrolysates with the stationary phase (Simpson, 2004). It also should be noted that upon hydrolysis the total integrated area of both SPIH and ESPIH increased markedly, suggesting increased protein solubility for both cases. Furthermore, it is noteworthy that at relatively low DH values (DH 5.3%), there was a considerable proportion of soluble protein with MW larger than 75.0 kDa (zone I) for ESPIH. For example, in ESPIH 5.3%, the proportion of soluble protein with MW larger than 75.0 kDa and in the range of 75.0e1.0 kDa (zone II) was 32.7% and 56.8%, respectively. In contrast, the corresponding proportion of soluble protein in ESPIH 9.1% was 11.7% and 77.3%, respectively. We inferred that the soluble protein components with MW larger than 75.0 kDa were mainly consisted of soluble protein aggregates. Our recent study have also found that limited enzymatic hydrolysis of
L. Chen et al. / Food Hydrocolloids 25 (2011) 887e897
SPI caused the release of the soluble protein aggregates from the insoluble protein aggregates (Luo et al., 2010).
3.3. Emulsifying properties of SPIH and ESPIH
3.3.1. SPI emulsions The mean droplet size of fresh emulsions formed by control SPI as a function of SPI concentration is shown in Fig. 6. As expected, mean droplet size decreased with the increase of SPI concentration. Because large emulsion droplets or droplet aggregates have higher weight in the calculation of d43 value than that in the calculation of d32 value, emulsions with similar d32 but different d43 (e.g., 2.0% w/v versus 2.5% w/v) would differ mainly in the amount of large droplets or droplet aggregates. At the SPI concentration of 5.0% (w/v), ﬁne oilin-water emulsions can be made with a mean droplet size of d32 z 0.41 mm and d43 z 1.48 mm. In this work, the concentration of SPI was chosen to be 1.6% (w/v) for making emulsions. Because the emulsion prepared using 1.6% (w/v) control SPI had a relatively large droplet size (d32 z 7.42 mm, d43 z 30.95 mm), there was scope for a substantial improvement on emulsifying capability of SPI using combined extrusion pre-treatment and enzymatic hydrolysis. 3.3.2. Inﬂuence of DH on emulsifying capability of SPIH and ESPIH The d43 of fresh emulsions formed by SPIH and ESPIH as a function of DH value is shown in Fig. 7. Emulsion formed by control ESPI had a large d43 of 27.43 mm. This result indicates that extrusion pre-treatment had a limited effect on improving the emulsifying capability of SPI. With the increase of DH value, d43 of both SPIH and ESPIH emulsion decreased initially, and then increased at high DH values. Hydrolyzed SPI showed a limited improvement, with a minimum d43 of 22.56 mm (SPIH 1.0%). In contrast, emulsions formed by ESPIH 8.0% or 9.1% exhibited huge improvement, with a d43 value as small as 2.01 mm. This result suggests that combined extrusion pre-treatment and enzymatic
Droplet size μ m
60 50 40 30 20 10 0 0
Concentration of SPI (w/v, %) Fig. 6. Inﬂuence of SPI concentration on mean droplet size of fresh emulsions (20% v/v oil, pH ¼ 7.0) formed by control SPI. Symbols: C, d32; B, d43.
60 The emulsifying properties of SPIH and ESPIH have been investigated via the formulation of 20% (v/v) sunﬂower seed oil-inwater emulsions at pH 7.0. Emulsifying capability refers to the ability of an emulsiﬁer to form and stabilize small droplets during homogenization, and was assessed in terms of mean droplet size and microstructure of fresh emulsions. Emulsion stability in this paper referred to the ability of an emulsion to resist creaming, ﬂocculation and coalescence over a storage period of 21 days, and was assessed by determining the changes in mean droplet size, microstructure and creaming index during storage.
45 30 15 0 0
DH (%) Fig. 7. Inﬂuence of DH on mean droplet size d43 of fresh emulsions (20% v/v oil, 1.6% w/ v emulsiﬁer, and pH ¼ 7.0) formed by SPIH and ESPIH. Symbols: ,, SPIH emulsions; A, ESPIH emulsions.
hydrolysis using pancreatin gives a much better effect on improving the emulsifying capability of SPI. 3.3.3. Microstructure observation of fresh SPIH and ESPIH emulsions Microstructures of some SPIH and ESPIH emulsions were visualized using CLSM. Droplet size distributions determined by light scattering (Mastersizer) were also superimposed on the microimages in order to highlight the correlations between particle size distribution and microstructure. Fig. 8AeC show the CLSM images and droplet size distributions of emulsions formed by SPIH with different DH values. As shown in Fig. 8A, emulsion formed by control SPI appeared to be highly ﬂocculated. The reason for this instability might be that there was insufﬁcient emulsiﬁer present at the newly created oilewater interface for a complete coverage. Therefore, bridging ﬂocculation of droplets became possible. Droplet size of control SPI emulsion was mainly distributed in the range of 10e100 mm. For the emulsion formed by SPIH 1.0% (Fig. 8B), the most interesting observation was the creation of some small droplets (ranged from 0.1 to 1 mm). However, ﬂocculation still occurred in this emulsion. This may indicate that SPIH 1.0% contained some peptides that were very surface active, enabling the formation of small droplets, but still the content of emulsiﬁer was not sufﬁcient for full coverage of droplet surface. For emulsion formed by SPIH 4.5% (Fig. 8C), droplet size distribution showed a shift to larger sizes in contrast with that of emulsions formed by control SPI and SPI 1.0%. Fig. 8DeF show the CLSM images and droplet size distributions for emulsions formed by ESPIH with different DH values. Emulsion formed by control ESPI contained a lot of big droplets and small droplet ﬂocs, and had a broad droplet size distribution ranging from 0.1 to 100 mm (Fig. 8D). Another interesting ﬁnding was that some elliptically shaped droplets were found in control ESPI emulsion. This may suggest that the interfacial membrane formed by control ESPI was easy to deform and less capable of protecting the droplets against coalescence. For emulsion formed by ESPIH 9.1% (Fig. 8E), the microstructure observation showed a ﬁne emulsion with a narrow droplet size distribution. And there was no sign of ﬂocculation for this emulsion. This observation indicated that emulsiﬁer present in ESPIH 9.1% was sufﬁcient to cover the entire newly created oilewater interface under the current conditions. Droplet size of ESPIH 9.1% emulsion was mainly distributed in the range of 0.1e10 mm. For emulsion formed by ESPIH 13.3% (Fig. 8F), both the
L. Chen et al. / Food Hydrocolloids 25 (2011) 887e897
Fig. 8. Microstructure of fresh emulsions (20% v/v oil, 1.6% w/v emulsiﬁer, and pH ¼ 7.0) formed by SPIH and ESPIH with selected DH values: (A) control SPI; (B) SPIH 1.0%; (C) SPIH 4.5%; (D) control ESPI; (E) ESPIH 9.1%; (F) ESPIH 13.3%. Droplet size distributions of diluted emulsions determined by light scattering (Mastersizer) are superimposed on the micrographs, with horizontal scale numbers indicating particle size in micrometers.
microstructure observation and the droplet size distribution measurement showed the appearance of some extraordinary large droplets. This suggests that excessive enzymatic hydrolysis caused a decrease in emulsifying capability for EPIH, as for SPIH. 3.3.4. Storage stability of emulsions formed by SPIH and ESPIH Storage stabilities of emulsions formed by SPIH (DH ¼ 0 or DH ¼ 1.0%) and ESPIH (DH ¼ 0 or DH ¼ 9.1%) were investigated over a storage period of 21 days with respect to creaming, ﬂocculation, and coalescence. Data of d43 and creaming index are
presented in Fig. 9A and B, respectively. Fig. 9A shows that the d43 of control ESPI emulsions increased signiﬁcantly after 21 days of quiescent storage, but it remained little changed for other emulsions. Fig. 9B shows that creaming was not observed in emulsion formed by ESPIH 9.1% after 21 days of quiescent storage, whereas it was signiﬁcant for other emulsions. These results suggest that emulsion formed by ESPIH 9.1% could be an ideal emulsiﬁer which was not only able to produce ﬁne emulsion droplets but also to provide excellent long term stability against ﬂocculation and creaming.
L. Chen et al. / Food Hydrocolloids 25 (2011) 887e897
1 day 21-day
d43 μ m
0 Control SPI
1 day 21-day
0 Control SPI
Fig. 9. Changes in properties of emulsions (20% v/v oil, 1.6% w/v emulsiﬁer, and pH ¼ 7.0) formed by SPIH (DH ¼ 0 or DH ¼ 1.0%) and ESPIH (DH ¼ 0 or DH ¼ 9.1%) during quiescent storage at ambient temperature for 21 days: A, mean droplet size d43; B, creaming index.
In order to explain the increase in d43 of control ESPI emulsion during storage, the changes in droplet size distribution and microstructure were examined. As shown in Fig. 10A, after stored for 21 days, the droplet size distribution of control ESPI emulsion showed a shift to larger size. A comparison of the CLSM image of the fresh emulsion (Fig. 8D) and that of the emulsion stored for 21 days (Fig. 10B) reveals that droplets coalescence occurred during storage.
Fig. 10. Changes in microstructure of emulsion (20% v/v oil, 1.6% w/v emulsiﬁer, and pH ¼ 7.0) formed by control ESPI during quiescent storage at ambient temperature for 21 days: A, droplet size distribution; B, CLSM image.
concentration increased, the interfacial pressure increased to a constant value (psat) when the interface became saturated with the emulsiﬁer. The measured values of psat were 16.2 mN/m, 15.2 mN/m, 14.3 mN/m and 10.1 mN/m for SPIH 1.0%, control SPI, ESPIH 9.1% and SPIH 4.5%, respectively. On the other hand, at low
3.4. Interfacial properties of SPIH and ESPIH with selected DH values π (mN/m)
As demonstrated before, combined extrusion pre-treatment and controlled enzymatic hydrolysis caused signiﬁcant changes in both physico-chemical properties and emulsifying properties for SPI. In order to better understand the relationship between them, some interfacial properties, including interfacial pressure (g) at sunﬂower seed oilewater interface, protein adsorption fraction during homogenization (Fads) and surface load (Gsat), were investigated for some SPIH and ESPIH with selected DH values.
12 9 6 3 0 1E-3
3.4.1. Interfacial pressure of SPIH and ESPIH at an oilewater interface The interfacial pressure at sunﬂower seed oilewater interface as a function of emulsiﬁer concentration for control SPI, SPIH 1.0%, SPIH 4.5% and ESPIH 9.1% is shown in Fig. 11. As the emulsiﬁer
Concentration of emulsifier (w/v, %) Fig. 11. Interfacial pressure at sunﬂower oilewater interface as a function of emulsiﬁer concentration for SPIH and ESPIH with selected DH values. Symbols: ,, control SPI; C, SPIH 1.0%; 6, SPIH 4.5%; A, ESPIH 9.1%.
L. Chen et al. / Food Hydrocolloids 25 (2011) 887e897
50 40 30 20 10 0
DH (%) Fig. 12. Inﬂuence of DH on protein adsorption fraction of SPIH and ESPIH during homogenization in the current condition (20% v/v oil, 1.6% w/v emulsiﬁer, and pH ¼ 7.0). Symbols: ,, SPIH emulsions; A, ESPIH emulsions.
protein concentrations less than 0.01% (w/v), the sequence of p values for different samples is SPIH 1.0% > control SPI > ESPIH 9.1% > SPIH 4.5%, which may indicate the sequence of their interfacial activity. Comparing interfacial activity with that of H0 for SPIH and ESPIH, we can see a positive correlation between the two parameters. 3.4.2. Fraction of protein adsorption and surface load of SPIH and ESPIH Fads of SPIH and ESPIH plotted as a function of DH value is shown in Fig. 12. Control SPI and control ESPI had a Fads of 31.6% and 28.2%, respectively. With the increase of DH value, Fads of both SPIH and ESPIH increased initially, and decreased at high DH values. ESPIH exhibited a maximum Fads of 67.6% at DH 9.1%, while SPIH showed a maximum Fads of only 38.9% at DH 1.0%. A comparison of Fig. 7 with Fig. 12 reveals a close relationship between the mean droplet size and the Fads for both SPIH and ESPIH. In most cases, the higher the Fads is, the smaller the d43 will be. Furthermore, it is interesting to note that ESPIH 5.3% and ESPIH 9.1% exhibited similar PS and H0 (see Table 1), but rather different Fads and d43. Fads of ESPIH 9.1% was larger than that of ESPIH 5.3%. And as expected, the d43 of ESPIH 9.1% emulsion should be smaller than that of ESPIH 5.3% emulsion. This difference may be caused by MWD factor. As has been demonstrated before, 77.3% of soluble proteins in ESPIH 9.1% were distributed in the MWD of 75.0e1.0 kDa, while only 56.8% of soluble proteins in ESPIH 5.3% were distributed in this MWD. However, ESPIH 5.3% contained a considerable proportion of soluble protein with MW larger than 75.0 kDa, which may consist of soluble protein aggregates. Therefore, we inferred that proteins or peptides in ESPIH with not too high a molecular weight may preferable for the adsorption to the oilewater interface during homogenization, as also reported by Ven, Gruppen, Bont, and Voragen (2001). Since controlled enzymatic hydrolysis can cause the unfolding of soy protein molecule, it could be possible that surface-active peptides with MWD of 75.0e1.0 kDa in ESPIH had a more ﬂexible microstructure. According to Dickinson (2009), biopolymers with ﬂexible structure are apt to lose their tertiary structure, and facilitate hydrophobic groups to direct towards the non-aqueous side of the interface, and therefore favour adsorption. The effectiveness of a protein as an emulsiﬁer also depends strongly on the surface load at saturation (Gsat). Emulsions formed
by 5.0% (w/v) control SPI and 1.6% (w/v) ESPIH 9.1% were chosen as representative systems for surface load studies. It was found that control SPI had a larger Gsat of 4.0 mg/m2 and ESPIH 9.1% had a smaller Gsat of 2.8 mg/m2. These Gsat values are comparable to that for most other food proteins which have been reported in the range of 1e10 mg/m2. Results of Gsat suggest that ESPIH 9.1% stabilized emulsions required less amount of protein than SPI stabilized emulsion. It has been suggested that ESPIH 9.1% may contain ﬂexible microstructure and therefore can rapidly alter their conformation, whereas rigid globular protein molecules have a slow conformation change due to kinetic constraints. The reason for this change has been explained by Bosker et al. (2003), who reckoned that biopolymers capable of undergoing fast conformational changes at the interface tended to have a lower surface load because of signiﬁcant spreading of the biopolymer molecules. To summarise, this work has studied the effects of combined extrusion pre-treatment and enzymatic hydrolysis on the interfacial properties and emulsifying capability of soy protein isolates. It was encouraging to ﬁnd that ESPIH 9.1% produced a very ﬁne emulsion (d32 ¼ 0.42 mm, d43 ¼ 2.01 mm) which exhibited excellent quiescent storage stability against droplet ﬂocculation and creaming. As compared with control SPI and SPIH 1.0%, ESPIH 9.1% appeared to be less surface active, mainly because of its lower H0. However, ESPIH has a slightly smaller psat but a much higher Fads (67.6%) and a signiﬁcantly lower Gsat (2.8 mg/m2). This may explain effective use of ESPIH in protecting emulsions against bridging ﬂocculation. As has been demonstrated, the enzymatic hydrolysis of ESPI was more intensive and extensive than that of SPI, because extrusion pre-treatment could signiﬁcantly improve the accessibility of SPI to enzymatic hydrolysis using pancreatin. Therefore, more soy proteins can be hydrolyzed and became soluble. In fact, extrusion pre-treatment and controlled enzymatic hydrolysis posed a striking effect on improving the solubility of SPI in the current study. The PS of ESPIH 9.1% was increased to 90.8%, much higher than that of SPI. Based on these results, we inferred that the strong increase in PS for ESPIH 9.1% probably was the main reason for the increase in its Fads. On the other hand, the decrease in MWD for ESPIH 9.1% probably caused an increase in ﬂexibility of protein structure, which was beneﬁted for the adsorption and unfolding of protein molecule at the oilewater interface. However, it should be noted that excessive enzymatic hydrolysis exert a negative effect on emulsifying capability for both SPIH and ESPIH. The reason for this result may be due to the similar changes in SPIH and ESPIH at high DH values: the strong decrease in H0. This could explain the decrease in both psat and Fads, and eventually resulted in the drastic decrease in their emulsifying capability. This study conﬁrmed that combined extrusion pre-treatment and controlled enzymatic hydrolysis using pancreatin led to signiﬁcant changes in physico-chemical properties and interfacial properties of SPI, and produced prominent beneﬁts in improving emulsifying capability of SPI and the stability of their emulsions. We demonstrated that combined extrusion pre-treatment and enzymatic hydrolysis could be an effective way for desirable functional modiﬁcation of various globular proteins.
Acknowledgements This research was supported by National Natural Science Foundation of China (Grant No. 20806030) and National High-tech R&D Program (863 Program) of China (Grant No. 2006AA10326). Thanks to Prof. E. Dickinson and Prof. B. Murray (University of Leeds) for useful discussion and advices. LC acknowledges ﬁnancial support from China Scholarship Council for his studies in the University of Leeds.
L. Chen et al. / Food Hydrocolloids 25 (2011) 887e897
References Achouri, A., & Zhang, W. (2001). Effect of succinylation on the physicochemical properties of soy protein hydrolysate. Food Research International, 34, 507e514. Adler-Nissen, J. (1986). Enzymatic hydrolysis of food proteins (427 pp.). London, U.K.: Applied Science Publishers. Akhtar, M., Dickinson, E., Mazoyer, J., & Langendorff, V. (2002). Emulsion stabilizing properties of depolymerised pectin. Food Hydrocolloids, 16, 249e256. Bosker, W. T. E., Agoston, K., Stuart, M. A. C., Norde, W., Timmermans, J. W., & Slaghek, T. M. (2003). Synthesis and interfacial behavior of polystyrenee polysaccharide diblock copolymers. Macromolecules, 36, 1982e1987. Burgaud, I., Dickinson, E., & Nelson, P. V. (1990). An improved high pressure homogenizer for making ﬁne emulsions on a small scale. International Journal of Food Science and Technology, 25, 39e46. Celus, I., Brijs, K., & Delcour, J. A. (2007). Enzymatic hydrolysis of brewers’ spent grain proteins and technofunctional properties of the resulting hydrolysates. Journal of Agricultural and Food Chemistry, 55, 8703e8710. Damodaran, S. (2004). Protein stabilization of emulsions and foams. Current Opinion in Colloid & Interface Science, 9, 305e313. Dickinson, E. (2009). Hydrocolloids as emulsiﬁers and emulsion stabilizers. Food Hydrocolloids, 23, 1473e1482. Govindaraju, K., & Srinivas, H. (2006). Studies on the effects of enzymatic hydrolysis on functional and physico-chemical properties of arachin. Food Science and Technology, 39, 54e62. Jung, S., Murphy, P. A., & Johnson, L. (2006). Physicochemical and functional properties of soy protein substrates modiﬁed by low levels of protease hydrolysis. Journal of Food Science, 70, 180e187. Kane, A. P., & Miller, D. D. (1984). In vitro estimation of the effects of selected proteins on iron bioavailability. American Journal of Clinical Nutrition, 39, 393. Kato, A., & Nakai, S. (1980). Hydrophobicity determined by a ﬂuorescence probe method and its correlation with surface properties of proteins. Biochimica et Biophysica Acta, 624, 13e20. Kinsella, J. E. (1979). Functional properties of soy proteins. Journal of the American Oil Chemists’ Society, 56, 242e258. Lee, K. H., Ryu, H. S., & Rhee, K. C. (2003). Protein solubility characteristics of commercial soy protein products. Journal of the American Oil Chemists’ Society, 80, 85e90. Liu, M., Lee, D. S., & Damodaran, S. (1999). Emulsifying properties of acidic subunits of soy 11S globulin. Journal of Agricultural and Food Chemistry, 47, 4970e4975. Luo, D. H., Zhao, Q. Z., Zhao, M. M., Yang, B., Long, X. T., Ren, J. X., et al. (2010). Effects of limited proteolysis and high-pressure homogenization on structural and functional characteristics of glycinin. Food Chemistry, 122, 25e30. Marsman, G. J. P., Gruppen, H., Mul, A. J., & Voragen, A. G. J. (1997). In vitro accessibility of untreated, toasted and extruded soybean meals for proteases and carbohydrates. Journal of Agricultural and Food Chemistry, 45, 4088e4095. McClements, D. J. (2005). Food emulsions: Principles, practice and techniques. Boca Raton, FL: CRC Press. Molina, E., Papadopoulou, A., & Ledward, D. A. (2001). Emulsifying properties of high pressure treated soy protein isolate and 7S and 11S globulins. Food Hydrocolloids, 15, 263e269. Nielsen, N. S. (1985a). The structure and complexity of the 11S polypeptides in soybeans. Journal of the American Oil Chemists’ Society, 62, 1680e1686.
Nielsen, N. S. (1985b). Structure of soy proteins. In A. M. Altschul, & H. L. Wilcke (Eds.), New protein foods (pp. 26e66). New York: Academic Press. Panyam, D., & Kilara, A. (1996). Enhancing the functionality of food proteins by enzymatic modiﬁcation. Trends in Food Science and Technology, 7, 120e125. Petruccelli, S., & Añón, M. C. (1994). Relationship between the method of obtention and the structural and functional properties of soy protein isolates. 1. Structural and hydration properties. Journal of Agricultural and Food Chemistry, 42, 2161e2169. Qi, M., Hettiarachchy, N. S., & Kalapathy, U. (1997). Solubility and emulsifying properties of soy protein isolates modiﬁed by pancreatin. Journal of Food Science, 62, 1110e1115. Radha, C., & Prakash, V. (2009). Structural and functional properties of heatprocessed soybean ﬂour: effect of proteolytic modiﬁcation. Food Science and Technology International, 15, 453e463. Roesch, R. R., & Corredig, M. (2003). Texture and microstructure of emulsions prepared with soy protein concentrate by high-pressure homogenization. Food Science and Technology, 36, 113e124. Rothenbuhler, E., & Kinsella, J. E. (1986). Disulﬁde reduction and molecular dissociation improves the proteolysis of soy glycinin by pancreatin in vitro. Journal of Food Science, 51, 1479e1482. Simpson, R. J. (2004). Purifying proteins for proteomics: A laboratory manual (pp. 201e202). New York: Cold Spring Harbor Press. Staswick, P. E., Hermodson, M. A., & Nielsen, N. C. (1984). Identiﬁcation of the cysteines which link the acidic and basic components of the glycinin subunits. Journal of Biological Chemistry, 259, 13431e13435. _ ski, D. (2004). Functional properties modiﬁcation of Surówka, K., & Zmudzi n extruded soy protein using neutrase. Czech Journal of Food Sciences, 22, 163e174. _ ski, D., Fik, M., Macura, R., & qasocha, W. (2004). New protein Surówka, K., Zmudzi n preparations from soy ﬂour obtained by limited enzymic hydrolysis of extrudates. Innovative Food Science and Emerging Technologies, 5, 225e234. Tsumura, K. (2009). Improvement of physicochemical properties of soybean proteins by enzymatic hydrolysis. Food Science and Technology Research, 15, 381e388. Tsumura, K., Saito, T., Kugimiya, W., & Inouye, K. (2004). Selective proteolysis of the glycinin and b-conglycinin fractions in a soy protein isolate by pepsin and papain with controlled pH and temperature. Journal of Food Science, 69, 363e367. Ven, C. V. D., Gruppen, H., Bont, D. B. A., & Voragen, A. G. J. (2001). Emulsion properties of casein and whey protein hydrolysates and the relation with other hydrolysate characteristics. Journal of Agricultural and Food Chemistry, 49, 5005e5012. Wagner, J. R., Sorgentini, D. A., & Añón, M. C. (2000). Relation between solubility and surface hydrophobicity as an indicator of modiﬁcations during preparation processes of commercial and laboratory-prepared soy protein isolates. Journal of Agricultural and Food Chemistry, 48, 3159e3165. Walstra, P. (1993). Principles of emulsion formation. Chemical Engineering Science, 48, 333e349. Walstra, P., & Smulders, I. (1997). Making emulsions and foams: an overview. In E. Dickinson, & B. Bergenstahl (Eds.), Food colloids: Proteins, lipids and polysaccharides (pp. 367e381). Cambridge: The Royal Society of Chemistry. Wu, W. U., Hettiarachcchy, N. S., & Qi, M. (1998). Hydrophobicity, solubility, and emulsifying properties of soy protein peptides prepared by papain modiﬁcation and ultraﬁltration. Journal of the American Oil Chemists’ Society, 75, 845e849.