Exploring the Relationship between Structural and Air–Water

Article pubs.acs.org/JAFC

Exploring the Relationship between Structural and Air−Water Interfacial Properties of Wheat (Triticum aestivum L.) Gluten Hydrolysates in a Food System Relevant pH Range Arno G. B. Wouters,*,† Ellen Fierens,† Ine Rombouts,† Kristof Brijs,† Iris J. Joye,†,§ and Jan A. Delcour† †

J. Agric. Food Chem. 2017.65:1263-1271. Downloaded from pubs.acs.org by RMIT UNIV on 01/08/19. For personal use only.

Laboratory of Food Chemistry and Biochemistry and Leuven Food Science and Nutrition Research Center (LFoRCe), KU Leuven, Kasteelpark Arenberg 20, B-3001 Leuven, Belgium § Food Science Department, University of Guelph, 50 Stone Road East Guelph, Ontario N1G 2W1, Canada ABSTRACT: The relationship between structural and foaming properties of two tryptic and two peptic wheat gluten hydrolysates was studied at different pH conditions. The impact of pH on foam stability (FS) of the samples heavily depended on the peptidase used and the degree of hydrolysis reached. Surface dilatational moduli were in most, but not all, instances related to FS, implying that, although the formation of a viscoelastic protein hydrolysate film is certainly important, this is not the only phenomenon that determines FS. In contrast to what might be expected, surface charge was not a major factor contributing to FS, except when close to the point-of-zero-charge. Surface hydrophobicity and intrinsic fluorescence measurements suggested that changes in protein conformation take place when the pH is varied, which can in turn influence foaming. Finally, hydrolyzed gluten proteins formed relatively large particles, suggesting that protein hydrolysate aggregation probably influences its foaming properties. KEYWORDS: gluten hydrolysates, pH, foam, air−water interface, protein conformation, interfacial properties

1. INTRODUCTION A foam is a dispersion of a gaseous phase in a liquid. Its creation requires energy input. Foams destabilize rapidly as they are thermodynamically unstable.1 Both foaming capacity (FC) and foam stability (FS) are important foam properties, which can be influenced by the presence of low molecular weight surfactants or proteins.2 Low molecular weight surfactants and proteins tend to diffuse to and adsorb at an air−water (A−W) interface. Once adsorbed, they stabilize the interface in different ways:1−3 • Proteins and low molecular weight surfactants at the interface lower the surface tension. • Proteins and ionic surfactants promote electrostatic repulsion between the gas bubbles they coat. • Proteins can sterically hinder gas bubbles from approaching each other and merging. • Proteins likely undergo changes in conformation upon adsorption at the A−W interface. They expose their more hydrophobic regions toward the air phase and form a viscoelastic film around the gas cells through protein− protein interactions, which stabilize the A−W interface. Although such interactions can be hydrogen bonds or of the van der Waals type, especially electrostatic and hydrophobic interactions are considered to be important in this context.1,2,4 Not all (plant) proteins are evenly suited as foaming agents. For example, wheat (Triticum aestivum L.) gluten protein, a coproduct of industrial starch isolation, has very low solubility in aqueous media and very low ability to form and stabilize foams.5−7 Enzymatic hydrolysis not only strongly increases the solubility of gluten but also improves its foaming properties.8 Such enzymatic hydrolysis induces three major structural © 2017 American Chemical Society

changes in proteins: a decrease in average molecular mass, better accessibility of hydrophobic regions, and a higher level of ionizable groups.9 These structural changes affect intermolecular interactions between the peptides in a hydrolysate and consequently their foaming and interfacial behaviors. Also, the structure of peptides as well as interpeptide interactions at an A−W interface are strongly influenced by environmental conditions such as pH. The effect of pH on plant protein hydrolysate foaming properties has been addressed by some researchers. The FC of pea protein isolates hydrolyzed with papain is lower at pH 8.0 than at pH 3.0, 5.0, and 7.0.10 A similar observation was made for peanut protein isolates hydrolyzed with Alcalase, which not only have good FC but also high FS at pH 3.0.11 In contrast, FS of barley protein hydrolysates (produced with subtilisin) increases with pH (up to pH 8.0), after which it decreases again (at pH 11.0).12 For gluten protein chymotrypsin hydrolysates, Popineau et al.13 reported better foaming properties (FC and FS) at pH 4.0 than at pH 6.5, whereas Drago and González14 observed an opposite trend for such hydrolysates when obtained using a fungal protease. They reported better foaming properties at pH 6.5 and 9.0 than at pH 4.0. Another study noted no differences in foaming capacity at pH 5.0, 7.0, and 8.0 of papain gluten hydrolysates.15 The impact of pH on foaming of wheat gluten hydrolysates is thus complex and depends on the peptidase used and the hydrolysis conditions. The latter determine which peptides are solubilized, the properties of which (including Received: Revised: Accepted: Published: 1263

November 11, 2016 January 17, 2017 January 26, 2017 January 26, 2017 DOI: 10.1021/acs.jafc.6b05062 J. Agric. Food Chem. 2017, 65, 1263−1271

Article

Journal of Agricultural and Food Chemistry their charge, conformation, and aggregation propensity) depend on conditions such as pH and temperature. For nonhydrolyzed proteins extracted from, for example, hemp seed,16 lentil,17 and soy,18 pH-induced structural changes and their effect on foaming have been described. For plant protein hydrolysates, similar studies relating pH to structural changes are rare. Some authors, including some of the ones mentioned above,10−12,14 have speculated on the molecular phenomena taking place at the A−W interface. However, it has often only been assumed but not experimentally shown [for example, by measuring zeta potential (ZP)] that the surface charge of proteins and peptides plays a key role in pH-dependent foaming.10,14 For example, Thewissen et al.19 observed pH- and salt-dependent foaming of gliadin hydrolysates from which they concluded this had to be related to the surface charge of the peptides. In addition to the above, pH-induced conformational changes in hydrolyzed plant protein that affect A−W interfacial properties remain to be investigated. This background shows that to better understand the structure−function relationship of wheat gluten hydrolysates, a comprehensive study is necessary. Here, foaming, A−W interfacial (decrease of surface tension and surface dilatational moduli), and structural (surface charge, protein conformation, and aggregation) properties of wheat gluten hydrolysates were evaluated at various pH values. The work reported has increased our understanding of how hydrolyzed proteins at A−W interfaces affect FS.

DH (%) =

X × Mx × 100 h = htotal α × M p × htotal

(1)

X is the consumption (mL) of NaOH or HCl solution needed to keep the pH during hydrolysis constant and Mx the molarity of the acid or base (respectively 0.50 and 0.20 M). The term α is a measure for the degree of dissociation of the α-NH3+ (neutral or alkaline conditions) or α-COOH group (acidic conditions). Under the given conditions, for tryptic hydrolysis α is 0.89,21 whereas for peptic hydrolysis it is 0.29.22 Mp is the mass of protein used, h is hydrolysis equivalents [milliequivalents (meqv)/g protein], and htotal is the theoretical number of peptide bonds per unit weight present in gluten protein. Nielsen et al.23 calculated the latter to be 8.3 meqv/g protein. 2.4. Measurement of Protein Solubility. First, solutions of T2, T6, P2, and P6 [0.075% (wprotein/v)] in 0.050 M sodium phosphate buffer (pH 6.8) containing 2.0% of sodium dodecyl sulfate (SDS) were made. Under such conditions, the samples were completely soluble and are further referred to as reference samples. Then, T2, T6, P2, and P6 [0.15% (wprotein/v)] were suspended in deionized water, and the pH was adjusted to pH 3.0, 5.0, and 7.0 by adding small amounts of 1.0 M NaOH or 1.0 M HCl. The samples were centrifuged (10 min, 10000g) to remove any insoluble material. An aliquot of the obtained supernatants (400 μL) was diluted with an extra 400 μL of 0.100 M sodium phosphate buffer (pH 6.8) containing 4.0% SDS. An aliquot (40 μL) of these diluted samples was loaded on a BioSep SEC-S3000 column (Phenomenex, Torrance, CA, USA) and analyzed using a Shimadzu (Kyoto, Japan) LC-2010 integrated HPLC system with peptide elution monitoring at 214 nm. Samples eluted at a flow rate of 0.50 mL/min using 0.050 M sodium phosphate buffer (pH 6.8) containing 2.0% SDS at 30 °C. Peptide solubility was determined by calculating the area in the chromatographic profiles and expressing it relative to the area in the profile of a reference sample (100% soluble). 2.5. Analysis of Foaming Properties. Foaming properties were determined with a standardized whipping test based on that of Caessens et al.24 Solutions of T2, T6, P2, and P6 [0.050% (wprotein/v)] were prepared, and the pH was adjusted to pH 3.0, 5.0, and 7.0 as above. An aliquot (50.0 mL) of these solutions was placed in a graduated glass cylinder (internal diameter = 60.0 mm) in a water bath at 20 °C. After temperature equilibration for 15 min, it was whipped for 70 s using a rotating propeller (outer diameter = 45.0 mm, thickness = 0.4 mm) at 2000 rpm. After whipping, the propeller was immediately removed and the glass cylinder sealed with Parafilm M (Bemis, Neenah, WI, USA) to avoid foam disruption by air circulation. The FC was the foam volume exactly 2 min after the start of whipping. The foam volume was then also measured at 4, 10, 15, 30, 45, and 60 min after the start of whipping. The decrease of foam volume over time was an indication for the FS of a given sample. On the basis of the foam height and the cylinder internal diameter, foam volume was calculated and expressed in milliliters. 2.6. (Oscillating) Pendant Drop Measurements. Solutions [0.050% (wprotein/v)] of T2, T6, P2, and P6 in deionized water, adjusted to pH 3.0, 5.0, and 7.0 as above, were introduced in a Theta optical tensiometer (Biolin Scientific Attension, Stockholm, Sweden) to create a pendant drop with a fixed volume of 8 μL. For every drop, the decrease in surface tension was measured over a 10 min time interval to assess protein adsorption and rearrangement at the A−W interface. In doing so, images were taken (1 frame per 7 s). A sinusoidal oscillation (50 cycles) was then performed at a frequency of 1 Hz with an amplitude set at 1.00 in the OneAttension software (Biolin Scientific Attension), which corresponded to a volume of ±1 μL. During oscillation, images were recorded (7 frames per second). The surface dilatational elastic modulus E′ of the interfacial film could be calculated from the drop shape analysis data during oscillation. Measurements were conducted at room temperature (22 ± 2 °C). After each measurement, the device was thoroughly cleaned and the surface tension of pure water was checked to be 72.0 ± 0.5 mN/m, before the next measurement was begun. 2.7. Measurements of Zeta Potential. Solutions of T2, P2, T6, and P6 [0.15% (wprotein/v)] in deionized water, adjusted to pH 3.0, 4.0, 5.0, 6.0, and 7.0 as above, were transferred to a disposable capillary

2. MATERIALS AND METHODS 2.1. Materials. Commercial wheat gluten was kindly provided by Tereos Syral (Aalst, Belgium). It contained 82.4% protein (N × 5.7) on a dry matter basis when determined using an adaptation of AOAC Official Method 990.0320 to an EA1108 Elemental Analyzer (Carlo Erba/Thermo Scientific, Waltham, MA, USA). Trypsin (EC 3.4.21.4) from porcine pancreas (reference from the distributor T0303) and pepsin (EC 3.4.23.1) from porcine gastric mucosa (reference from the distributor P6887) were purchased from Sigma-Aldrich (Bornem, Belgium), as were all other chemicals, solvents, and reagents. 2.2. Enzymatic Hydrolysis. A 6.0% (wprotein/v) wheat gluten aqueous dispersion was incubated with trypsin or pepsin at pH-stat conditions in a Titrino 718 device (Metrohm, Herisau, Switzerland). For each enzyme, gluten was hydrolyzed to degrees of hydrolysis (DH) of 2 and 6. The DH reflects the percentage of initially present peptide bonds that have been hydrolyzed (see below). For tryptic hydrolysis, pH-stat conditions were 50 °C and pH 8.0, and an enzyme to substrate ratio of 1:480 (DH 2) or 1:20 (DH 6) on a protein mass basis was used. For peptic hydrolysis, the reactions were carried out at 37 °C and pH 3.5, and an enzyme to substrate ratio of 1:1200 (DH 2) or 1:300 (DH 6) on pa rotein mass basis was used. When the desired DH was reached, the pH was adjusted to 6.0 and proteolysis was stopped by heating the protein suspension for 15 min at 95 °C. It should be noted that the heating procedure may cause limited peptide aggregation. This will be further discussed under section 3.6. The mixtures were then centrifuged (10 min, 12000g) at room temperature, and the supernatants were filtered and then freeze-dried. All further analyses, including those of protein contents (carried out as outlined under section 2.1), were conducted on the freeze-dried supernatants of DH 2 or DH 6 tryptic (further referred to as T2 and T6, respectively) and peptic (further referred to as P2 and P6, respectively) hydrolysates. 2.3. Determination of Degree of Hydrolysis. DH is the percentage of peptide bonds hydrolyzed (h) relative to the total number of peptide bonds (htotal) per unit weight present in wheat gluten protein. It was calculated from the quantity of NaOH (trypsin) or HCl (pepsin) solution used to keep the pH constant during hydrolysis: 1264

DOI: 10.1021/acs.jafc.6b05062 J. Agric. Food Chem. 2017, 65, 1263−1271

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Journal of Agricultural and Food Chemistry zeta cell (Malvern Instruments, Malvern, UK) to determine the ZP in a Zetasizer Nano ZS (Malvern) based on laser Doppler microelectrophoresis. Measurements were conducted at room temperature (22 ± 2 °C). 2.8. Determination of Surface Hydrophobicity. The surface hydrophobicity of all samples was determined with 1-anilino-8naphthalenesulfonic acid (ANS) as fluorescent probe. Samples were dissolved in deionized water, and the pH was adjusted to pH 3.0, 4.0, 5.0, 6.0, and 7.0 as above. They were then diluted with deionized water (adjusted to the same pH) to protein concentrations of 0.18, 0.36, 0.54, 0.72, and 0.90 mg/mL. Of each diluted sample, 200.0 μL was transferred to a 96-well plate in duplicate, and 10.0 μL of ANS solution (8.0 mM in deionized water) was added. The fluorescence emission intensity of the protein samples was recorded at 480 nm with a Synergy Multi-Mode Microplate Reader (BioTek, Winooski, VT, USA) after excitation at 390 nm. The relative fluorescence intensity was calculated as the intensity of the fluorescence of the protein−ANS mixture minus that of the control sample (ANS with water) fluorescence, which was then divided by the fluorescence intensity of the control sample. The slope of the plot of relative fluorescence intensity as a function of protein concentration for each sample represents the surface hydrophobicity. Measurements were conducted at room temperature (22 ± 2 °C). 2.9. Analysis of Intrinsic Tryptophan Fluorescence. Changes in the local environment of tryptophan residues in a protein chain can be assessed from the fluorescence spectrum of the tryptophan indole ring. The intrinsic tryptophan fluorescence of T2, T6, P2, and P6 solutions [0.010% (wprotein/v) in deionized water] adjusted to pH 3.0, 5.0, and 7.0 as above was evaluated with a Fluoromax 4 fluorospectrometer (Horiba Jobin Yvon, Edison, NJ, USA). The excitation wavelength was 295 nm with a 2 nm slit width, whereas the emission spectrum was recorded from 300 to 450 nm using a 3 nm slit width. Measurements were conducted at room temperature (22 ± 2 °C). 2.10. Dynamic Light Scattering. Solutions of T2, P2, T6, and P6 [0.150% (wprotein/v) in deionized water] adjusted to pH 3.0, 5.0, and 7.0 as above were placed in disposable cuvettes to measure dynamic light scattering in a Zetasizer Nano ZS (Malvern). The particle size distribution (PSD) was calculated from this scattering pattern with the Stokes−Einstein relationship using the Malvern Zetasizer software. Measurements were conducted at room temperature (22 ± 2 °C). 2.11. Statistical Analysis. Protein solubility of the samples at different pH values was determined in duplicate. All foam-related experiments, ZP, dynamic light scattering, surface hydrophobicity, pendant drop, and intrinsic tryptophan fluorescence measurements were carried out at least in 4-fold. All data were analyzed using statistical software JMP Pro 11 (SAS Institute, Cary, NC, USA), with a Student t test at a significance level α = 0.05.

letters) except for P2, which had a marginally lower solubility at pH 5.0 and 7.0 than at pH 3.0. In addition, at pH 5.0 there were no significant (p < 0.05) differences in solubility between the four different samples. At pH 7.0, P6 had a slightly lower protein solubility than the other samples, whereas at pH 3.0, it was slightly less soluble than P2. Thus, none of the differences observed in foaming or structural properties described in the following paragraphs can be attributed to differences in peptide solubility. The reason that only minor differences were observed is that all insoluble material had already been removed directly after hydrolysis, whereas in most other studies on the impact of pH on plant protein hydrolysates (see the Introduction) this was not the case.10,12−15 We believe that the merit of our approach is that it allows drawing conclusions about the relationship between gluten hydrolysate structural properties and their foaming, thereby surpassing mere differences in levels of solubilized protein. 3.2. Foaming Properties. Foaming properties of the four hydrolysates were affected differently by pH (Figure 1). At a given pH value, the FC did not differ much between different samples (Figure 1). The FC of all samples was significantly (p < 0.05) higher at pH 7.0 than at pH 3.0 or pH 5.0, except for T2, which had higher FC at pH 7.0 than at pH 3.0 but not than that at pH 5.0. Overall, these differences were minor. In terms of FS, all samples sustained at least some foam after 60 min at pH 7.0. However, DH 2 hydrolysates clearly had better FS than DH 6 hydrolysates. At pH 5.0, foams from all hydrolysates destabilized very quickly as illustrated by the steep, fast decrease in foam volume. At pH 3.0, tryptic and peptic hydrolysates clearly behaved differently. Whereas both foams of peptic hydrolysates very rapidly collapsed at pH 3.0, those of tryptic hydrolysates did not. The foam of T2 at pH 3.0 was the most stable of all foam samples produced. Although the foam volume of T6 at pH 3.0 after 60 min was zero, it did not rapidly lose stability the way foams of P2 and P6 did. Thus, a pH decrease from 7.0 to 5.0 led to a steep decrease in FS for all hydrolysates. A further pH drop from 5.0 to 3.0 improved FS of tryptic but not peptic hydrolysates. These differences in FS, not only for different samples at a given pH but also for a given sample at different pH values, might be attributed to the nature of protein films formed at the interface at different pH values. Electrostatic and hydrophobic interactions, because of differences in the charge of ionizable groups and peptide conformation at various pH values, probably play a role in this context. All of these hypotheses will be explored in the following paragraphs. 3.3. Air−Water Interfacial Properties. The foaming of protein hydrolysates depends on their adsorption at the A−W interface and the nature of the film they there form. Differences in pH values may alter the surface activity and the ability of gluten hydrolysates to form a strong viscoelastic protein film. Figure 2 shows the decrease of surface tension over time after formation of a pendant drop. The rate and extent of this decrease are indicative of the rates of diffusion and adsorption of the hydrolysate constituents at the interface, as well as of their rearrangement at the A−W interface, thereby orienting their more hydrophobic regions toward the air phase. For both T2 and T6, the decreases in surface tension were similar at pH 3.0, 5.0, and 7.0 (Figure 2), so the rate and ability of their peptides to diffuse to and continuously adsorb at the A−W interface remained unchanged. Proteins continue to adsorb and rearrange at the interface until an equilibrium is reached. Data were collected up to 10 min after drop formation. At this time,

3. RESULTS AND DISCUSSION 3.1. Protein Solubility. The solubility of the peptides ranged from 94 to 100% (Table 1). No impact of pH on solubility was noted for any of the samples (Table 1, capital Table 1. Impact of pH on the Protein Solubility of T2, T6, P2, and P6 Hydrolysatesa pH 3.0 T2 T6 P2 P6

99 97 101 95

± ± ± ±

3 1 1 1

A,ab A,ab A,a A,b

pH 5.0 95 94 97 96

± ± ± ±

2 2 1 1

A,a A,a B,a A,a

pH 7.0 99 98 98 94

± ± ± ±

0 1 0 1

A,a A,a B,a A,b

a Capital letters indicate significant (p < 0.05) differences between different pH values for a given sample. Lower case letters indicate significant (p < 0.05) differences between samples at a given pH value. The codes T and P refer to gluten trypsin and pepsin hydrolysates; the codes 2 and 6 refer to degrees of hydrolysis of 2 and 6.

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Figure 1. Impact of pH on the foaming properties of 0.050% (wprotein/v) solutions of T2, T6, P2, and P6 hydrolysates. The codes T and P refer to gluten trypsin and pepsin hydrolysates; the codes 2 and 6 refer to degrees of hydrolysis of 2 and 6.

both tryptic hydrolysates gradually decreased, whereas the opposite was observed for both peptic hydrolysates. These had the highest E′ values at pH 7.0. As already stated above, foams of both DH 2 hydrolysates at pH 7.0 were more stable than those of both DH 6 hydrolysates. This is in accordance with the fact that T2 and P2 had significantly (P < 0.05) higher E′ than T6 and P6 at pH 7.0. However, not all observations regarding FS were fully in line with E′ values. For example, all samples had relatively high E′ at pH 5.0, even though their foams rapidly lost stability at this pH. This suggests that the elasticity obtained through oscillating drop measurements of the protein film in some, but not all, cases, and especially not at pH 5.0, can explain differences in FS. 3.4. Surface Charge. As pH evidently affects the charge of the ionizable groups in the hydrolysates and thereby may alter their ability to interact at the A−W interface and to stabilize the foam, it may well be at the basis of differences in foaming. At pH 7.0, all samples had comparable ZPs (p > 0.05), with values ranging from −2 to −4 mV, except for T6, which had a much lower ZP (−16 mV) (Figure 3). This observation cannot directly be related to differences in foaming properties. Indeed, although P6 and T6 had similar low FS at pH 7.0, the former had a much higher ZP than the latter. In addition, these two DH 6 hydrolysates had comparable surface tensions (Table 2) and surface dilatational moduli (Figure 2) at pH 7.0. As expected, for all hydrolysates, the zeta potential increased with decreasing pH, as the ionizable groups were gradually protonated. The point-of-zero-charge (PZC), that is, the pH at which a sample has a ZP of zero, of T2, P2, and P6 was about 5.0, whereas that recorded for T6 was about 4.0. At this pH,

the surface tension approached a plateau, and it was possible to estimate a value representative of the equilibrium surface tension (Figure 2). To this end, surface tension versus 1/√t (t = time) plots (not shown here) were made using the data in Figure 2. From a certain time onward (and thus at low 1/√t values) linear relationships were observed, which were used to make extrapolations to 1/√t = 0 (which accords to t → ∞). The latter yielded surface tension values, indicative of equilibrium surface tensions (Table 2). Even though there were some significant differences in these estimated equilibrium surface tension values of either T2 or T6 at different pH values (Table 2, capital letters), these differences were minor. In contrast, peptic hydrolysates were much less able to decrease surface tension at pH 3.0 than at pH 5.0 and 7.0. Also, estimated equilibrium surface tension values for P2 and P6 were much higher at pH 3.0 than at pH 5.0 or 7.0 (Table 2, capital letters), suggesting that the hydrolysates had a lower affinity for the A−W interface or that they were not able to arrange themselves very efficiently at the A−W interface at this lower pH value. These observations are in line with the fact that both tryptic hydrolysates led to a reasonable FS at pH 3.0, whereas peptic hydrolysates did not. After subsequent sinusoidal volume oscillations of the hanging drops, surface dilatational moduli could be determined (Figure 2). E′, which is indicative of the elasticity of the film at the interface, is often related to foam stability, in the sense that elevated film elasticities in many instances are responsible for high stability in protein foams.3,25 T2 and, to a lesser extent, T6 had very high E′ at pH 3.0. As expected from the low FS of peptic hydrolysates at pH 3.0, P2 and P6 had very low E′ at pH 3.0. With increasing pH, E′ of 1266

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Figure 2. Impact of pH on the decrease of surface tension at the air−water (A−W) interface after forming a hanging drop with 0.050% (wprotein/v) solutions of T2, T6, P2, and P6 hydrolysates. After 10 min, the drops were sinusoidally oscillated to determine surface dilatational moduli (E′). Codes T2, T6, P2, and P6 are as in Figure 1.

Table 2. Estimation of Equilibrium Surface Tension of 0.050% (wprotein/v) Solutions of T2, T6, P2, and P6, at pH 3.0, 5.0, and 7.0, at t → ∞ Based on Surface Tension versus 1/√t Plots Using the Same Data As Displayed in Figure 2a pH 3.0 T2 T6 P2 P6

48.6 50.8 54.4 58.7

± ± ± ±

0.5 0.4 1.3 0.7

pH 5 B,d A,c A,b A,a

49.0 50.0 49.7 52.1

± ± ± ±

1.0 0.7 0.3 0.6

pH 7 B,c B,b B,bc B,a

51.1 50.4 49.5 51.8

± ± ± ±

0.6 0.5 0.5 0.9

A,ab AB,b B,c B,a

a Capital letters indicate significant (p < 0.05) differences for a given sample at different pH values. Lower case letters indicate significant (p < 0.05) differences between samples at a given pH value. Codes T2, T6, P2, and P6 are as in Table 1.

electrostatic repulsion between peptide chains is minimal, so aggregationwith or without loss of solubilityis most probable. These observations are largely in line with the very low FS at pH 5.0 of all samples. However, as discussed under section 3.1, there was no decrease in protein solubility at pH 5.0. It is indeed possible that aggregation occurs at the PZC but apparently only to such an extent that it does not affect peptide solubility. Furthermore, despite the low FS readings at pH 5.0, all hydrolysates still had relatively high E′ values. Thus, the lack of an overall net charge did not prevent the hydrolysates from interacting and forming rather strong protein films. At pH 3.0, although there were some statistically significant (P < 0.05) differences in ZP between some of the samples, these should not be overinterpreted as all ZP values were rather close to zero

Figure 3. Impact of pH on the zeta potential (ZP) of 0.150% (wprotein/ v) solutions of T2, T6, P2, and P6 hydrolysates. Codes T2, T6, P2, and P6 are as in Figure 1.

and differences were only minor. In contrast, there were large differences in FS as well as E′ values between tryptic and peptic hydrolysates at pH 3.0. Thus, the assumption that the pH dependence of foaming is dictated by the charge of ionizable groups in the hydrolysates does not hold true in all cases. Only at the PZC, where FS is very low for all samples, does surface charge seem to be directly related to differences in foaming. We 1267

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the carboxylic groups originating from the backbone of the protein chain have a pKa of around 2.0 and for a significant part will not occur in their protonated form at pH 3.0.29 Additionally, if this were the sole factor involved, it would have been expected that all hydrolysates would have had a similar response, as they had comparable ZPs at pH 3.0. Clearly, this is not the case. Thus, there are some arguments which suggest that the peptides in the hydrolysates might in fact undergo pH-induced conformational changes. These could then contribute to the pH dependence of their foaming, especially because the surface hydrophobicity increased drastically at pH 3.0, which is the pH at which tryptic and peptic hydrolysates had distinctly different foam stabilities. However, as similar ZP values (and therefore net surface charges) do not always correspond to equal levels of ionizable groups, such possible interference by electrostatic interactions between ANS and the peptides at pH 3.0 could not be completely ruled out. Therefore, changes in protein conformation were also studied by analysis of tryptophan fluorescence. Changes in the local environment of this amino acid affect both the wavelength at which the emission intensity is highest (i.e., the maximum emission wavelength) and the fluorescence emission spectrum intensity. Shifts in maximum emission wavelength usually indicate changes in polarity of the environment of the amino acid, whereas changes in intensity are often related to a higher mobility and accessibility of tryptophan for quenchers. Thus, both phenomena report on changes in protein conformation.30 Figure 5 shows the impact of pH on the emission spectra (300−450 nm) of all four hydrolysates. There were notable shifts in the maximum emission wavelengths neither for different samples at a given pH nor within one sample at different pH values. Values of maximum emission wavelengths all ranged between 355 and 359 nm, which is a value typical for tryptophan residues that are completely exposed to a polar environment. At the wavelength of maximum emission and at any given pH value, P2 had the highest fluorescence intensity, followed by T2, T6, and P6, respectively (all differences were significant, p < 0.05). Additionally, for any given sample, the fluorescence emission intensities at pH 7.0 were significantly (p < 0.05) higher than at pH 5.0, which in turn were significantly higher than the values recorded at pH 3.0. All of this confirmed that it is likely that changing the pH induces changes in the conformation of the proteins. Furthermore, as all samples had a similar change in fluorescence emission intensity, this and the surface hydrophobicity values at different pH values did not explain why tryptic hydrolysates had relatively high FS, whereas peptic hydrolysates had very low FS, at pH 3.0. 3.6. Particle Size Distribution. In the previous paragraphs, it was suggested that the peptides may occur as aggregates. Even when it would not affect the overall peptide solubility, such aggregation may heavily influence the A−W interfacial properties of gluten hydrolysates. PSDs were measured with dynamic light scattering. There was quite some variation between different measurements of the same sample at a given pH. This may be due to the fact that aggregates present in such hydrolysates are loosely assembled and reorganize rather easily upon changes in the environment. The PSDs shown in Figure 6 are averages of at least 12 repeated measurements and are indicative of the aggregation of different samples under these conditions. In all cases, samples contained particles ranging from 10 to 4000 nm in size (Figure 6). Thus, enzymatic hydrolysis probably yielded peptides that have a tendency to

hypothesize that the low FS observed at pH 5.0 (and thus close to the PZC) is likely related to a lack of electrostatic repulsion between different gas bubbles, which makes their coalescence more likely, rather than to a weakening of the protein films around the gas bubbles. This would mean that at pH 3.0 and 7.0, repulsion (because of the overall positive or negative charge of the protein films) occurs to such an extent that it prevents gas bubbles from easily merging. At these pH values, the strength and integrity of the protein films (see section 3.3) probably determine the stability of gluten hydrolysate foams. 3.5. Surface Hydrophobicity and Intrinsic Tryptophan Fluorescence. As the pH changes, not only the charge of the peptides varies but also their conformation may be affected. For various native plant proteins, pH-induced conformational changes have been studied,16−18 but this has not yet been reported for plant protein hydrolysates. In the present case, irrespective of pH, peptic hydrolysates had a higher surface hydrophobicity than tryptic hydrolysates, and DH 2 hydrolysates had a higher surface hydrophobicity than their DH 6 counterparts (Figure 4). These results are in accordance with

Figure 4. Impact of pH on the surface hydrophobicity of solutions of T2, T6, P2, and P6 hydrolysates. Codes T2, T6, P2, and P6 are as in Figure 1.

RP-HPLC evidence from our team26 that at pH 6.4, P2 and P6 have higher overall hydrophobicity than T2 and T6. The highest surface hydrophobicity was observed at pH 3.0 for all samples. Compared to the surface hydrophobicity at pH 4.0, an 8-fold increase of surface hydrophobicity was observed for T2 at pH 3.0, followed by P6 (5.5-fold increase), P2 (5-fold increase), and finally T6 (2-fold increase). These surface hydrophobicity readings (Figure 4) at pH 3.0 seem to suggest that hydrophobic regions are more accessible by ANS at this pH value, possibly indicative of strong differences in conformation. With regard to the surface hydrophobicity readings, it should be mentioned that due to the negative charge on the sulfonic acid group of ANS, electrostatic interactions between proteins and ANS may influence surface hydrophobicity measurements.27,28 Whereas, upon the transition from pH 4.0 to pH 3.0, there are barely any functional groups that become positively charged, there is an increase in net positive charge of the peptide mixture due to the protonation of negatively charged carboxylic acid groups. However, although the carboxylic groups of the side chains of glutamic acid and aspartic acid have pKa values of around 4.0, 1268

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Figure 5. Impact of pH on the intrinsic tryptophan fluorescence emission spectra of solutions of T2, T6, P2, and P6 hydrolysates after excitation at 295 nm. Codes T2, T6, P2, and P6 are as in Figure 1.

Figure 6. Impact of pH on the particle size distribution (PSD), measured with dynamic light scattering of 0.150% (wprotein/v) solutions of T2, T6, P2, and P6 hydrolysates. Codes T2, T6, P2, and P6 are as in Figure 1.

with care. PSDs of P2 and P6 were not or only slightly affected by variations in the pH. It seemed that, although intrinsic fluorescence and, even more so, surface hydrophobicity

form (soluble) aggregates. It should be mentioned that larger particles scatter more light, meaning that their relative abundance based on the PSDs in Figure 6 should be interpreted 1269

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in protein conformation took place in gluten hydrolysates when the pH was varied. Such changes seemed most pronounced when the pH was changed to 3.0, which is the pH value at which striking differences in FS between peptic and tryptic hydrolysates were observed. However, at this pH, there was a clear distinction neither in surface hydrophobicity nor in fluorescence intensity between tryptic and peptic hydrolysates. It is, of course, possible that different conformational rearrangements lead to a similar response in fluorescence intensity, but with a completely different effect on interfacial stabilization. At the same time, the techniques used here allow only the evaluation of conformational changes of the hydrolysates in solution, whereas peptide behavior at the A− W interface might still be different. Furthermore, it is important to consider protein hydrolysate aggregation, as it became clear that hydrolyzed gluten formed relatively large but probably loose aggregates, which may influence foaming. Such aggregates can to a certain extent be considered “particles”. Upon adsorption at an A−W interface, inorganic particles provide excellent stability against disproportionation and coalescence by forming a strong mechanical barrier.37 However, research on biodegradable, food-grade, protein-based particles has only recently emerged, and much is still to be learned.38−40 More research is needed to assess the role of peptide aggregation on the A−W interfacial and foaming behavior of protein hydrolysates. Finally, we conclude that all measurements performed here were necessary to better understand how structural properties of plant protein hydrolysates change with pH and how this is related to their foaming behavior. Still, not all observations could be explained, illustrating the complexity of studying the behavior of proteins at the A−W interface.

measurements indicated that conformational changes occurred in P2 and P6, these changes did not affect their aggregation, as measured by dynamic light scattering. In contrast, the impact of pH on the PSDs of tryptic hydrolysates was much more pronounced. T2 had polydisperse and rather similar PSDs at pH 3.0 and 7.0. Whereas at pH 7.0 T2 and T6 both had polydisperse PSDs, the latter had a higher average particle size. At pH 3.0, a monodisperse peak of particles at around 530 nm was observed for T6. At pH 5.0, all samples had rather monodisperse PSDs. One could hypothesize that the aggregates at pH 5.0 would be larger than those at other pH values because of the proximity to the PZC, where no electrostatic repulsion occurs, thereby leading to formation of an impaired protein film at the A−W interface. However, this was not always the case. For example, T6 solutions contained even larger (>1000 nm) particles at pH 3.0 than at pH 5.0, but led to better FS at pH 3.0 than at pH 5.0. It is difficult to assess how the presence of these aggregates affects the A−W interfacial and foaming properties of gluten hydrolysates. Aggregation may slow the diffusion to and adsorption at the interface but may also affect the ability of the hydrolysate constituents to rearrange themselves at the A−W interface. Rullier et al.31,32 showed that β-lactoglobulins are more efficient at forming foams in a nonaggregated state than when present in a heatinduced aggregated form. In contrast, the presence of aggregated proteins led to higher thin film and foam stability. Such effects have also been described for food proteins such as those of pea,33 soy,34 egg white,35 and whey.35,36 Thus, the presence of aggregated as well as nonaggregated peptides in gluten hydrolysates may contribute to their overall foaming characteristics. From the results in Figure 6, no direct relationship between the aggregation state of the hydrolysates at different pH values and their A−W interfacial behavior (see section 3.3) could be established. However, given the relatively large variation of the particle size distributions (as described above), it is likely that the aggregates observed here are not very compact and that they rearrange or dissociate in the whipping procedure to produce foams. A more systematic study on the role of controlled aggregation of protein hydrolysates on the way they stabilize A−W interfaces is desirable. In conclusion, depending on the enzyme used for their production and the degree of hydrolysis reached, different gluten hydrolysates were affected in very different ways by changes in pH. At pH 7.0, DH 2 hydrolysates had better FS than DH 6 hydrolysates, whereas at pH 3.0, tryptic hydrolysates had better FS than peptic hydrolysates. At pH 5.0, which was close to the PZC of the hydrolysates, they all had relatively high E′ values, but very low FS. We believe this is due to a lack of electrostatic repulsion between gas bubbles, making their coalescence more likely, irrespective of the strength of the protein films. At pH 3.0 and 7.0 the hydrolysates had an overall positive or negative charge, so there was some repulsive effect between gas bubbles. At these pH values, protein film elasticity values were in line with their FS. Thus, surface charge was a major factor that directly contributed to FS in the proximity of the PZC. At other pH values, FS depended on the strength and integrity of the protein films at the A−W interface, to which hydrophobic interactions probably contributed significantly. Although pH-induced changes in protein conformation for native (plant) proteins have been studied in various instances, prior to the present work this was not the case for protein hydrolysates. Surface hydrophobicity and intrinsic fluorescence measurements performed here did in fact suggest that changes

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AUTHOR INFORMATION

Corresponding Author

*(A.G.B.W.) Phone: +32 (0) 16 372035. E-mail: arno. [email protected]. ORCID

Arno G. B. Wouters: 0000-0002-6868-4432 Funding

I.R. thanks the Research Foundation − Flanders (FWO, Brussels, Belgium) for financial support. K.B. acknowledges the Industrial Research Fund (KU Leuven, Leuven, Belgium) for a position as Industrial Research Manager. J.A.D. is W. K. Kellogg Chair in Cereal Science and Nutrition at KU Leuven. This work is part of the Methusalem program “Food for the Future” at KU Leuven. Notes

The authors declare no competing financial interest.

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ABBREVIATIONS USED FC, foaming cacapity; FS, foam stability; A−W, air−water; ZP, zeta potential; DH, degree of hydrolysis; h, number of hydrolyzed peptide bonds; htotal, total number of peptide bonds present; Mp, mass of protein; Mx, molarity of acid or base; meqv, milliequivalents; SDS, sodium dodecyl sulfate; ANS, 1-anilino-8-naphthalenesulfonic acid; PSD, particle size distribution; t, time; PZC, point-of-zero-charge

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