The sulfhydryl group and disulfide bond content were determined using Ellman’s reagent according to the method described by Pan et al. (23). Elman’s reagent was obtained by dissolving 40 mg of 5,5′-dithiobenzoic acid (DTNB) with 10 ml Tris–Gly buffer solution (1.2 g of disodium ethylene diamine tetraacetic acid [EDTA], 6.9 g of Gly, and 10.4 g of Tris in 1,000 ml of distilled water; pH 8.0). Lyophilized samples obtained after different treatments were dissolved in Tris-Gly-SDS buffer (90 ml Tris-Gly buffer containing 10 ml of SDS aqueous solution), bringing the final protein concentration to 10 mg/ml. The samples were lysed and centrifuged at 8,000 × g for 20 min, following which 4 ml of the supernatant was mixed with 0.04 ml of Ellman’s reagent and incubated in the dark for 30 min. Finally, the absorbance was measured at 412 nm using a spectrophotometer to determine the free sulfhydryl content. To examine the total sulfhydryl content, lyophilized protein samples were dissolved in a urea-Tris-glycine buffer containing (10 M urea, 2.5% SDS, 0.004 M EDTA, 0.09 M glycine, and 0.086 M Tris; pH 8.0), mixed evenly, and incubated for 30 min. Subsequently, the samples were centrifuged at 8,000 × g for 20 min, and the supernatant was dissolved in 5 ml of 12% TCA and centrifuged again. This wash step was repeated 3 times. The precipitate was collected and dissolved in 2.5 ml of Tris-glycine buffer, followed by the addition of 50 ml DTNB and incubation for 30 min under dark conditions. The overall content of sulfhydryl groups was detected based on the absorbance at 412 nm. The molar extinction factor was 1360 M^–1cm^–1. Additionally, the content of the disulfide bonds was obtained by subtracting the free sulfhydryl level from the total sulfhydryl level and dividing it by 2. The levels of sulfhydryl groups and disulfide bonds were calculated as follows:

where D is the coefficient of dilution; A₄₁₂ indicates the absorbance of QA in Ellman’s reagent; C_t is the content of total sulfhydryl groups; and C_f is the content of the free sulfhydryl groups.

The solubilities of the samples were analyzed as described by Abugoch and Romero (25). In brief, the prepared 1% w/v QA suspensions were magnetically stirred for 1 h and centrifuged at 7,500 × g for 20 min. Bradford’s method was used for measuring the amount of QA in the supernatant. Solubility was indicated as a percentage of the total albumin.

The water (WBC) and oil binding capacity (OBC) of QA were measured based on the methods used by Beuchat (26). The WBC was calculated as the grams of water retained per gram of sample, and the OBC was measured as the amount of oil held per gram of sample.

Emulsifying activity (EA) and emulsion stability (ES) assays were conducted according to the methods used by Zhang et al. (27). First, 15 ml of QA solution (10 mg/ml) was mixed with 5 ml corn oil using a homogenizer at 23,000 rpm for 2 min. Then, 2 ml of the homogenized sample was mixed with deionized water (18 ml), from which 0.5 ml was removed and mixed with 4.5 ml of 0.1% SDS. The absorption was measured at 500 nm using 0.1% SDS as the control:

where A₀ is the sample absorbance value; DF represents the fold dilution (100); C is the emulsion solution that had a concentration before use (g/ml); φ is the oil volume fraction; θ is the oil phase fraction of 0.25; and A₃₀ represents the absorbance at 30 min.

Foaming capacity (FC) and foam stability (FS) measurements were adapted based on the protocol described by Aydemir and Yemenicioglu (28). First, 20 ml of 1% QA solution obtained after different treatments were homogenized at 23,000 rpm for 2 min. The mixture was poured into a 50-ml measuring cylinder, and the volume was recorded. The FC and FS were calculated according to the following formulae:

where V₀ is the volume prior to homogenization (ml) and V₁₀ is the volume after homogenization (ml).

Simulated gastric fluid (SGF) and simulated intestinal fluid (SIF) were prepared according to the protocol detailed by Minekus et al. (29). In brief, SGF was prepared by mixing 0.7 ml HCl (12 M) with 2.0 g NaCl and 0.3 g pepsin to a final volume of 100 ml (pH 1.2). SIF was prepared by completely dissolving 0.68 g KH₂PO₄ in 25 ml deionized water, and later adding 19 ml NaOH (0.2 M) and 40 ml deionized water, and finally 4.0 g trypsin (pH 7.5).

In vitro digestion was performed as described by Cho (30). QA solution (5 mg/ml) and an equivalent amount of SGF were placed in a centrifuge tube and shaken in a 37°C water bath for 30, 60, 90, and 120 min. The digests were immediately boiled at 100°C (10 min) to kill the enzymes and then cooled quickly on the ice. After adjusting the pH of the remaining solution to 7.5, an equivalent amount of SIF was added and shaken at 37°C. The digestion products were removed at 150, 180, 210, 240, 270, and 300 min, boiled at 100°C, and then cooled and set aside.

O-phthalaldehyde (OPA) was used to determine the degree of QA hydrolysis (31). For this test, 400 μl of hydrolyzed supernatant was mixed with 3 ml OPA, and after 2 min of incubation, the absorbance at 340 nm was measured. A serine standard solution (0.9516 mM) and deionized water were employed as the standard and blank, respectively. To calculate the degree of hydrolysis, the following formulae was used:

where SerineNH₂ is the amount of Ser per each g protein (mmol/g); X is the sample quality (g); p is the sample protein content (%); N is the number of dilutions; and V is the supernatant volume (L).

where α and β represent the constants 1 and 0.4, respectively, and the h_tot of QA is 7.4 mmol/g.

The digestion products were lyophilized, and the method for AA analysis was identical to the AA analysis method described previously (section “Amino acid”).

Sulfhydryl groups function as precursors of disulfide bonds, which are extremely important for the tertiary structure of a protein. Typically, an increase in sulfhydryl content indicates alterations to a protein’s structure (37). Therefore, sulfhydryl and disulfide bond analysis were indispensable for evaluating the structural and functional changes in QA (38). As shown in Figure 2, with increasing temperature, the content of sulfhydryl groups first increased and then decreased (P < 0.05). Moreover, the opposite trend was observed for disulfide bonds (P < 0.05). The highest content of sulfhydryl groups (2.16 nmol/mg) and lowest content of disulfide bonds (3.18 nmol/mg) were found at 80°C. The above data indicated that moderate heating (50–80°C) could increase the sulfhydryl group content while reducing the disulfide bonds in QA. This could be due to the unfolding of protein structure under heat treatment and thus the subsequent exposure of interior sulfhydryl groups. When protein subunits dissociate, disulfide bonds are broken and sulfhydryl is generated (39). Nevertheless, the opposite trend was the case under too high-temperature treatment that QA may undergo partial dissociation of protein aggregates, exposing more disulfide bonds. The changes in SH of heated-QA were similar to the irradiated quinoa albumin. The SH content of rice albumin increased significantly as the irradiation dose, this tendency could be ascribed to its higher sulfur-containing amino acids (i.e., methionine) and more disulfide bonds destructed by irradiated (40). Combining the result of increased SH content, it was supposed that disulfide bonds of proteins were destroyed under heat treatment and then exposed to the solution, which contributed to the solubilization.

Hydrophobicity indicates the degree of exposure to surface hydrophobic groups and is closely associated with the interfacial activity of a protein (41). As seen in Figure 3, heat treatment had a significant effect on the H₀ of QA. Specifically, H₀ values initially decreased with increasing temperatures and then increased thereafter (P < 0.05), reaching the lowest value of 4.28 at 80°C (approximately 57% of the control). Notably, the comparable trend was detected for H₀ changes of flaxseed albumin at different pH conditions, this phenomenon could be attributed to the variations in molecular interaction in the protein molecules, leading to denaturation of protein molecules (42). This lower hydrophobicity could be due to the fewer charges on the protein surface, which causes protein aggregation, thus burying hydrophobic sites in the aggregated structures following heat treatment (43). In contrast, when the temperature is too high, protein molecules can unfold, exposing buried hydrophobic groups and resulting in increased hydrophobicity (44). Likewise, similar results were obtained when the hydrophobicity of protein isolates from heat-treated legumes was compared with that from the untreated group (45).

Solubility is a measure of the degree of protein aggregation and denaturation, as well as an indicator of protein functionality (46). As depicted in Figure 4, the solubility of QA gradually increased with increasing temperature (P < 0.05), achieving a peak of 95% at 60°C. It then declined progressively as the temperature continued to rise (P < 0.05). Protein solubility is determined by the interactions between protein and water molecules, on the one hand, and the aggregated properties of protein fraction in the process of heating, on the other hand (47). Hence, the appropriate conditions of heating can promote the hydration capacity of a protein and further increase its solubility. Yet, once the temperature becomes too high, the protein denatures, the conformation of the protein molecule changes, and the hydrophobic groups present in the protein are exposed, leading to reduced protein solubility. This trend is similar to that observed from the solubility of the chicken bone protein in ultrasonic power study, possibly due to the cavitation and thermal effects that increase the energy of molecules in the system (48).

The WBC and OBC of a protein are crucial metrics to be considered while formulating food products (49). The mouthfeel and texture of foods are affected by the WBC, and the OBC is associated with the emulsion properties of products such as mayonnaise (50). As shown in Figure 5, the strength of water binding in QA increased progressively with a rise in temperature, but after 60°C, a gradually decreasing trend was observed (P < 0.05). Variations in WBC may be caused by the thermally induced unwinding of protein segments that occurs when the temperature increases, and there is an increase in the hydrophilic groups on the surface, enhancing the interaction force. On the contrary, when the temperature rises beyond a certain threshold, the aggregation of denatured proteins may lead to a decrease in their ability to bind to water. The OBC of QA tended to first rise and then fall with increasing temperature (P < 0.05), with the maximum value being reached at 60°C. This was probably due to the near-complete unfolding of the proteins during the process of heating, which increases the interaction between the small molecules and the oil, thereby expanding the OBC. Nevertheless, the oil’s viscosity decreases, and fluidity increases with a further increase in temperature, resulting in a weaker OBC. Similar types of results for WBC and OBC were observed in heat-treated sunflower protein isolates (44). The improvement in WBC and OBC were also noted in nano fibrillated whey protein isolates after the application of heat treatment (51). Thus, changes in protein conformation with increasing binding sites under appropriate heating could lead to better WBC and OBC functionality (52). Heat treatment provides fresh ideas for included baked goods, cereals, ice creams, various dessert formulations, and chocolates.

Generally, EA functions to measure the capacity of interfacial adsorption onto the oil-water interface of proteins, and ES is employed to examine the protein’s ability to be retained on an oil-water interface after storage in an emulsion (53). To evaluate the emulsion properties of QA, its EA and ES were measured in this study. As shown in Figure 6, a similar trend was observed in EA and ES variation. Both values first increased with temperature, peaking at 60°C, and then tended to decrease thereafter (P < 0.05). There is evidence of a tight relationship between the solubility and emulsification properties of a protein (54). In the present study, the emulsification ability of QA was markedly affected by heat treatment, probably as a result of the increased solubility, which allowed the rapid spreading and adsorption of QA at the oil-water interface (46) and thus elevating its emulsification capacity (55). A similar kind of observation was obtained for the emulsion activity and stabilities of heat-treated vanillin-rich protein isolates from kidney beans (45). The emulsifying activities of proteins are affected by their molar mass, hydrophobicity, conformational stability, charge and physicochemical factors such as pH, ionic strength, and temperature (56), in which, solubility is regarded as a prime factor for the emulsifying properties of protein isolates. Steaming and baking exert negative impacts on the emulsion stability of quinoa protein isolate, which might be related to the aggregation of protein molecules and the reduction of viscosity (57).

Foaming capacity represents the size of the interfacial amount produced by a protein during the process of foaming, while FS represents the ability of a protein to stabilize bubbles under the influence of gravity (44). The FC and FS of QA after different heat treatments are presented in Figure 7. A gradual and continuous elevation of FC was observed with an increase in treatment temperatures (P < 0.05), but this value began to plateau after 70°C (P > 0.05). During the process of heating, protein chains are partially unraveled. This allows them to be more easily absorbed at the air-water interface, thus increasing FC (58). Additionally, FS exhibited an initial decrease, reaching its minimum at 70°C, and then tended to increase with increasing temperatures subsequently. The improvements in FS may be the result of increased protein-protein interaction (aggregation) during heating, leading to the formation of a thick protein membrane surrounding the produced bubbles (59). Ding et al. (60) found that the trend of FC under other physical treatments was similar to that of heat treatment, which showed an upward trend with increasing intensity of physical conditions, followed by a downward trend. At the same time, we found that the heated albumins with over 85% of FC which was significantly higher than the corn germ meal albumin of FC (67%) (61).

An in vitro digestion assay was conducted by simulating artificial gastrointestinal fluid. Changes in the hydrolysis rate represent the degree of peptide bond cleavage during simulated digestion (62). As illustrated in Figure 8, the degree of hydrolysis of QA increased with prolongation of digestion time (P < 0.05). Moreover, heat-treated groups (80, 100, and 121°C) showed lower degrees of hydrolysis than the control, with a similar tendency across the process of digestion. This is compatible with findings from Tian et al. (63) which showed that the degree of hydrolysis of soy protein isolates reduces after thermal treatment above 85°C. Similarly, Sangsawad et al. (64) also reported that high-temperature treatment at 121°C strongly decreases the digestibility of chicken breast protein. The change in QA digestibility after heat treatment can be attributed to cross-linked protein aggregates, which exhibit good resistance to hydrolysis (65). However, these results were found to be the opposite of those of germinated legume proteins after soaking treatment (66). High-pressure processing treated pea protein underwent a greater degree of proteolysis and showed different peptide patterns after static gastric digestion compared to untreated and heat-treated pea protein (67).

Changes in the total AA content of QA during digestion following different treatments are shown in Figure 9, and the classification of the AAs can be seen in Table 2. The total AA levels at the end of trypsin digestion (300 min) were remarkably higher than those at the end of pepsin digestion (120 min) irrespective of treatment temperature (P < 0.05). Moreover, the content of AAs in the in vitro digestion product of unheated QA was higher than that in heat-treated samples (P < 0.05). Among them, the AA content after heat treatment at 121°C was the lowest (11.70 mg/g) at the end of digestion, 45.1% lower than the control. These results are consistent with those from Shi et al. (68) which demonstrated that heat treatment significantly reduces the AA content of apricot mushroom proteins following in vitro digestion. Similar findings have also been reported regarding lower AA levels following the in vitro digestion of bran proteins after microwave heat treatment (69). The extent of protein digestion in the gastrointestinal tract can be indicated by the levels of Arg, Lys, Phe, Trp, and Tyr, which are the targeted cutting sites for pepsin and pancreatic proteases (70). Our experimental results support the findings that heating reduces the content of Arg, Lys, Phe, Trp, and Tyr in QA (P < 0.05). As a result, we speculate that high-temperature thermal treatment may cause severe destruction to AAs and induce irreversible decomposition, leading to a loss of contents and finally a reduced hydrolysis ratio. Joycelyn et al. (71) found that heat-induced protein unfolding led to a 38% decrease in the amount of Cys in amino acid compared to untreated cowpea proteins, whereas combined thermal-ultrasonic treatment increased Cys content of the hydrolysates by 41%. This will be an interesting subject for future QA processing that heating technology could combine with non-heat ones to enhance digestibility.