Time-Dependent Effects of Ultrasonic Modification of Soy Protein Concentrate on the Mixolab Rheology of Enriched Dough

Abstract

Soy protein concentrate (SPC) often has limited food applications due to the loss of its functional properties under harsh industrial processing. This study explored the effects of exposure time to high-intensity ultrasound (HUS) on the structural properties of SPC to assess the potential of a single protein for multiple bakery applications. HUS treatment modified SPC free sulfhydryl group content (4.81 ± 0.03 to 1.47 ± 0.01 µmol/g_protein) and hydrophobicity (34.17 ± 0.02 to 30.56 ± 0.03 µgBPB/mg_protein) and promoted the formation of soluble and insoluble aggregates, especially with longer exposure times, as evidenced by SDS-PAGE. According to Raman analysis, SPC exposed to 0.5 min HUS exhibited an α-helical content of 33.52 ± 1.58% and β-sheet content of 56.80 ± 4.40%, while the tyrosine doublet (I₈₅₀/I₈₃₀) ratio was associated with dough stability and indicated intermolecular hydrogen bonding within the dough matrix. Water absorption capacity was improved upon addition of HUS-exposed SPC samples, to 58.4 ± 0.71%, compared with 52.6 ± 0.85% of SPC-enriched dough. These changes accelerated dough development time and enhanced amylase activity, resulting in a dough with desirable viscosity. HUS-exposed samples with higher α-helix content and solubility, decreased water syneresis, and hydrophobic SPC formed stabile complexes with hydrophobic regions of the amylose chain, both leading to reduced starch retrogradation (1.551 ± 0.13 to 0.855 ± 0.04). Overall, this study showed that by controlling the HUS treatment time, protein structure can be tailored for its use in diverse bakery applications, further enhancing the commercial value of protein concentrates.

1. Introduction

Soy protein concentrate (SPC) is a by-product obtained during soybean oil production, following the removal of residual oil using hexane and the soluble carbohydrate fraction [1]. Owing to its high protein content (60–68%) and good digestibility, SPC is considered a viable plant-based alternative to animal proteins in food systems. Namely, these proteins have good nutritional value due to their adequate amino acid composition; techno-functional properties, such as oil- and water-binding capacity; and emulsifying properties, giving them wide application [2].

Despite the associated health-promoting effects of soy proteins, consumption of soy products in Western European countries remains below recommended levels (only 1.5% of men and 2.1% of women reported soy consumption) [3]. Incorporation of soy proteins into bread and bakery products, thus making them a part of the daily diet, represents a viable strategy for increasing overall soy protein intake in Western Europe. However, wheat flour fortification with soybean flour and protein, in order to obtain quality products in terms of taste, consumer acceptability and shelf life, remains a real challenge. Although fortification improves the overall protein content and balances the essential amino acid composition of wheat flour, it also impairs the rheological and sensory properties of the final product, which is reflected in darker color, low bread loaf volume, reduction in biscuit width, and increase in biscuit thickness [4]. In addition, the bread’s hardness and density increases, as does the viscosity of the dough, but the main impediment to soy flour application is actually the beany flavor, derived from lipoxygenase activity [5]. Therefore, there is a clear need to improve the color, functional, and sensory properties of soy protein products (soy flour, SPC, soy protein isolate (SPI)) to enhance consumer acceptance and increase the intake of soy protein–supplemented bakery products. The negative effects associated with soy–wheat dough appear to be due to the lack of interactions between soy and gluten proteins, which depends on several factors, including the content of functional groups (thiol groups, hydrophobic and charged residues capable of forming electrostatic interactions) on their surfaces, enabling them to interact with each other [6]. Although dough is a complex food matrix containing other ingredients, and thus, the mode of interaction between soy proteins and wheat proteins in dough is still unclear, several researchers have confirmed that the hydrophobic interactions are important for the stabilization of these associated molecules [6,7].

High-intensity ultrasound (HUS; 20–100 kHz) holds its appeal as a highly efficient and widely applicable technology across various materials. It is an environmentally friendly, noninvasive technology proven in its potential to reshape the structure of proteins, making them tailorable for specific food applications [8]. During HUS treatment, the acoustic waves pass through the medium and stretch and compress the structure in the medium, forming tiny bubbles in each stretching phase. As the process of stretching and compressing continues, these bubbles collapse and grow. These bubbles can violently explode or not, meaning there are two types of cavitation: transient and stable [9,10]. Acoustic cavitation accelerates protein unfolding, denaturation, and aggregation. Which of these effects will be dominant depends on the type of protein, duration of treatment, temperature increase, intensity (W/cm³), energy/density (W/mL; J/mL), size and shape of the sonoreactor, and solvent ratio [11]. In general, ultrasonic cavitation and shear forces induce changes in proteins’ secondary and tertiary structures by breaking hydrophobic and hydrogen bonds, resulting in conformational changes [11]. Ultrasonic treatment has been widely applied to modify the structure and functional properties of various proteins, including whey protein, rapeseed, chickpea, egg white, and pea protein, resulting in changes in solubility, emulsifying capacity, foaming and gelation properties, as well as antioxidant activity [12,13,14,15,16,17]. There are numerous data about HUS application in the functionalization of SPI, which testify to a decrease in particle size and increase in solubility and hydrophobicity, and, consequently, emulsifying and foaming capacity [18,19,20,21]. On the other hand, data on the impact of HUS on SPC are scarce [19,22]. Considering that SPC is a more effective and industrially relevant alternative to SPI, further investigation of its functional modification by HUS is scientifically and practically justified.

This study was aimed at linking the main structural and functional changes induced by HUS in SPC with the rheological properties of wheat flour formulation containing 5% w/w of the corresponding SPC. Structural changes were monitored and analyzed through measurements of surface hydrophobicity and the content of disulfide groups, while changes in the secondary structure were enlightened by Raman spectroscopy. The Mixolab instrument was used to explore the relationship between rheological changes and structural modifications in SPC induced by HUS by measuring fundamental parameters, including dough absorption, dough development time, dough stability, protein resistance to heat and mechanical mixing, starch gelatinization rate, amylase activity, as well as starch retrogradation. This work presents new insights into how ultrasonic modification of the structural and functional properties of soy protein concentrate affects dough rheology, which has not been previously reported in the literature.

2. Materials and Methods

2.1. Materials

Soy protein concentrate (SPC, TRADCON F-200) was used as the primary material in this study. It contained 70.96 ± 1.77% dry weight basis (N × 6.25) protein, with a moisture content of 5.81%, ash content of 6.41 ± 0.51%, fat content of 0.06 ± 0.03%, cellulose content of 3.80 ± 0.18%, and total fiber content of 20.63 ± 1.44%. This product is manufactured by SOJAPROTEIN, located in Bečej, Serbia.

The 5,5′-dithiobis-(2-nitrobenzoic acid) (DTNB) was obtained from Sigma-Aldrich Co. in St. Louis, MO, USA. Ethylenediaminetetraacetic acid disodium salt dihydrate (EDTA) was sourced from Tokyo Chemical Industry UK Ltd., Oxford, UK. Additionally, commercial T-500 wheat flour (Triticum aestivum) was acquired from the supplier Sentella in Senta, Serbia and used as reference material for the rheological tests conducted in this study. The proximate analysis of the flour indicated a maximum moisture content of 15%, protein content of 10.48% (dry weight basis), ash content of 0.45% (dry weight basis), starch content of 69.39% (dry weight basis), and fat content of 0.98% (dry weight basis). All other chemical reagents used were of analytical grade.

2.2. Methods

2.2.1. Sample Preparation and HUS Pretreatment

Soybean protein concentrate suspension (8%, w/v) was prepared by homogenizing the sample in deionized water in a 100 mL batch reactor. For ultrasonic treatment, a Bandelin Sonopuls HD 2200 ultrasonic homogenizer equipped with a titanium sonotrode probe TT13 (Ø 13 mm), Sonopuls HD 2200 (Bandelin Electronic GmbH & Co. KG, Berlin, Germany) was used. The ultrasonic probe assembly, which consists of converter UW 2200, booster horn SH 213 G and TT13 titanium sonotrode, was placed inside a Bandelin noise protection box (Bandelin Electronic GmbH & Co. KG, Berlin, Germany) in order to reduce noise emission levels and stabilize operating conditions. Samples were treated for 0.5, 2.0, and 5.0 min at a constant amplitude of 30% and a frequency of 20 ± 0.2 kHz, with temperature controlled using an ice–water bath. During treatment, the active tip of the TT13 sonotrode, only a few millimeters in length, remained fully submerged in the suspension, while the non-active upper part stayed above the liquid level, ensuring stable transfer of ultrasonic energy into the system. Subsequently, the samples were spray-dried using a Mini Spray Dryer (BUCHI Labortechnik AG, Flawil, Switzerland) under the following conditions: inlet temperature 160 °C, outlet temperature approximately 70 °C, aspirator 100%, and feed flow rate 50%, controlled using the flow rate valve.

2.2.2. Evaluation of Protein Surface Properties via Sulfhydryl Group Determination

The content of free and total sulfhydryl groups was determined using the Ellman reagent, specifically, 5,5′-dithio-bis-2-nitrobenzoic acid (DTNB) [23]. A protein dispersion was prepared to assess the free sulfhydryl groups by mixing the protein concentrate in sodium phosphate buffer (0.1 M, pH 8.0) with 1 mM EDTA. For the total sulfhydryl groups, the protein concentrate was dispersed in the same sodium phosphate buffer but with the addition of 2% (w/v) SDS. In both cases, the protein concentrates were mixed in a solid–liquid ratio of 2.5:1 with corresponding phosphate buffer containing DTNB at a concentration of 0.3 g/L. The resulting dispersions were incubated in the dark at room temperature for 15 min, with constant stirring. After incubation, the samples were centrifuged at 6700× g for 5 min at room temperature (MiniSpin^®, Eppendorf AG, Hamburg, Germany), and the absorbance of the supernatant was measured at 412 nm against buffer. The content of free or total sulfhydryl groups was calculated using the molar extinction coefficient of 13,600 M⁻¹ cm⁻¹, and the results were expressed as micromoles of free or total sulfhydryl groups per gram of protein (µmol/g protein):

$$Sulfhydryl group content \left({\displaystyle \frac{\mathsf{\mu }\mathrm{m}\mathrm{o}\mathrm{l}}{{\mathrm{g}}_{\mathrm{p}\mathrm{r}\mathrm{o}\mathrm{t}\mathrm{e}\mathrm{i}\mathrm{n}}}}\right)={\displaystyle \frac{∆A_{412\mathrm{n}\mathrm{m}}\times d_f}{\epsilon \times l\times c_{protein}}}$$

(1)

where ∆A_412nm is the absorbance of the sample at 412 nm; d_f is the dilution factor; ε = 13,600 M⁻¹ cm⁻¹ is the molar extinction coefficient of DTNB; l is the path length (cm); and c_protein is the protein concentration (g/L).

2.2.3. Evaluation of Protein Surface Characteristics via Surface Hydrophobicity Determination

Surface hydrophobicity was assessed using the bromophenol blue dye (BPB) fixation method [24]. The sample (5.0 mg) was suspended in 50 mM phosphate buffer at pH 7.0. The dye was dissolved in a small volume of ethanol sufficient for complete solubilization, and distilled water was added to reach a final concentration of 1.0 mg/mL. To analyze the samples, 0.2 mL of the BPB dye solution was added to 1.0 mL of the sample suspension. A control sample consisting of 1.0 mL of buffer and 0.2 mL of dye solution was prepared simultaneously. The samples were incubated at room temperature with constant stirring for 15 min. Following the incubation, the samples were centrifuged at 9650× g for 10 min (MiniSpin^®, Eppendorf AG, Hamburg, Germany). The supernatant was then diluted with buffer in a 1:10 ratio, and the absorbance was measured at 595 nm relative to the buffer. The obtained absorbance values were subsequently used for calculations according to the following equation:

$$\mathrm{B}\mathrm{P}\mathrm{B} \left({\displaystyle \frac{\mathsf{\mu }\mathrm{g}}{{\mathrm{m}\mathrm{g}}_{\mathrm{p}\mathrm{r}\mathrm{o}\mathrm{t}\mathrm{e}\mathrm{i}\mathrm{n}}}}\right)={\displaystyle \frac{200 \mathsf{\mu }\mathrm{g}}{m_{protein}}}\times \left({\displaystyle \frac{A_{control}-A_{sample}}{A_{control}}}\right)$$

(2)

where A_control is the absorbance of the BPB dye solution mixed with buffer (without protein) and A_sample is the absorbance of the BPB dye solution mixed with the protein sample suspension after incubation and centrifugation, and m_protein is the mass of protein (mg).

2.2.4. SDS-PAGE Profiling of HUS-Modified SPC

SDS-PAGE analysis was performed using an mPAGE™ 10 × 8 cm, 12% Bis-Tris gel according to the Laemmli method [25]. Prior to electrophoresis, 10.0 mg of each sample was suspended in 0.5 mL of distilled water and incubated at room temperature for 30 min under continuous mixing. The suspensions were then mixed in a 1:1 ratio with sample buffer—either reducing (prepared with 2.5 mL of Tris-HCl buffer, pH 6.8, 2.0 mL glycerol, 2.0 mL of 1 M DTT solution, 4.0 mL of 10% SDS, and 5.0 mg of bromophenol blue) or non-reducing (same buffer without DTT). After mixing, the samples for reducing SDS-PAGE were incubated in a boiling water bath for 5 min and then centrifuged at 2400× g for 5 min (Thermo Fisher Scientific, Waltham, MA, USA), whereas samples for non-reducing SDS-PAGE were loaded onto the gel immediately after mixing with the buffer and centrifugation. Electrophoresis was conducted using a vertical Hoefer S260 Mighty Small II Mini Vertical Electrophoresis System with 1.5 mm combs/spacers (Hoefer Inc., Holliston, MA, USA). A volume of 6.0 µL from each prepared sample was loaded into individual wells, and protein separation was carried out for 70 min at 120 V in mPAGE MOPS SDS running buffer. Following electrophoresis, the gel was stained with Coomassie Brilliant Blue solution (prepared with 100.0 mg of dye, 40% methanol, and 10.0% glacial acetic acid) for 15 min. The gel was then destained until protein bands became clearly visible.

2.2.5. Characterization of Molecular Protein Structure via Raman Spectroscopy

The effect of HUS treatment on the SPC molecular structure was analyzed by Raman spectroscopy, whereby the Raman spectra were recorded using a Raman spectrometer (XploRA, Horiba Jobin Yvon, Kyoto, Japan) at room temperature. The laser excitation wavelength was 532 nm (maximum output power 20–25 mW). Spectra were collected using an acquisition time of 15 s and a boxcar smoothing set of 2. All measurements were realized using the spectrometer equipped with 1800 g mm⁻¹ grating. The Raman spectra (200–1800 cm⁻¹) of each sample were performed in triplicate, and the results were reported as the average of these replicates with a relative standard deviation of less than 5%. Immediately before quantitative analysis, the obtained spectra were normalized to the intensity of the phenylalanine band at 1004 cm⁻¹, which was used as an internal standard to compensate for variations in signals between samples. Normalized values of characteristic regions of the Raman spectrum, such as tyrosyl doublet, phenylalanine band, S-S stretching band and δ-CH band, were isolated. The conformational composition of the disulfide bonds was analyzed in the S-S region located at 490–550 cm⁻¹. This spectral region was extracted and deconvoluted by Gaussian fitting. Based on the literature, three conformational forms were detected: the gauche–gauche–gauche (g–g–g, interchain) conformation with peaks in the range of 500–515 cm⁻¹, the gauche–gauche–trans (g–g–t, intrachain) conformation appearing at 520–530 cm⁻¹, and the trans–gauche–trans (t–g–t, intrachain) conformation located at 540–550 cm⁻¹ [26]. The composition of the secondary structure was determined from the amide I region located in the range of 1600–1700 cm⁻¹. Before analysis, smoothing was performed using the Savitzky–Golay algorithm with a second-order polynomial and a 5-point window to reduce noise using OriginPro^® 2022 software. Then, the amide I band was deconvoluted using a Gaussian function, and the peak positions were determined by the second derivative method. The positions of the secondary structure elements were adopted from the literature, with β-sheet structures at 1610–1640 cm⁻¹, random coil at 1640–1650 cm⁻¹, α-helical structures at 1650–1660 cm⁻¹ and β-turns at 1660–1690 cm⁻¹ [27]. The relative content of each secondary structure component was calculated from the area under the corresponding fitted peaks.

2.2.6. Determination of Thermo-Mechanical Properties of Dough Partially Substituted with HUS-Modified SPC

The thermo-mechanical properties of wheat flour dough and wheat flour dough enriched with different SPC types were analyzed using the Mixolab analyzer (Chopin Technologies, Villeneuve-la-Garenne, France) in accordance with the “Chopin +” protocol [28]. The mixtures tested with the Mixolab were prepared externally by mixing wheat flour with the corresponding protein concentrate to obtain 5% (w/w) mixture. During the simulation of the mixing and baking process in the Mixolab bowl, the sample was subjected to heating and cooling depending on the analysis phase, and the torsion (Nm) was measured. At the end of the analysis (45 min), the following data was determined from the Mixolab curve: water absorption (WA), dough development time (DDT), and dough stability. The characteristic peak values of the Mixolab curve are C1 → 1.05 ± 0.05 Nm (desired consistency); C2 → protein weakening as a function of mechanical work and temperature; C3 → starch gelatinization; C4 → hot gel stability; and C5 → starch retrogradation during the cooling phase.

2.2.7. Statistical Analysis

Statistical analysis of the data was performed using OriginPro^® 2022 software (OriginLab Corporation, Northampton, MA, USA). All experiments were conducted in triplicate, and results are presented as mean ± standard deviation. To evaluate the statistical significance of the differences in means, a one-way analysis of variance (ANOVA) was conducted, followed by Tukey’s test to confirm any significant differences. Differences were considered statistically significant at p < 0.05.

3. Results and Discussion

3.1. Influence of HUS Treatment Time on Sulfhydryl Group Content and Surface Hydrophobicity

Sulfhydryl groups are highly reactive functional groups found within protein structures. They can undergo oxidation and form both intra- and intermolecular disulfide bridges. These bridges help stabilize proteins’ tertiary structures, which in turn shape their functional properties [29]. Similarly, during dough mixing, sulfhydryl groups are oxidized, forming intra- (within the protein) and intermolecular (between proteins) disulfide bonds, reflecting changes in dough formation and stability [30]. Following the specific aim of this manuscript to modify the rheological properties of dough with structurally modified soy proteins, we examined the effect of the ultrasonic exposure time on the content of total and free sulfhydryl groups (

Table 1. Impact of exposure time to HUS probe on free/total sulfhydryl group content and surface hydrophobicity of SPC (amplitude—30% and frequency—20 ± 0.2 kHz).

Sample Label	Free Sulfhydryl (µmol/gprotein)	Total Sulfhydryl (µmol/gprotein)	Hydrophobicity (µgBPB/mgprotein)
SPC	4.81 ± 0.03 a	7.91 ± 0.02 a	34.17 ± 0.02 a
SPC-0.5	1.02 ± 0.03 d	1.74 ± 0.02 d	23.41 ± 0.06 d
SPC-2	1.27 ± 0.02 c	2.06 ± 0.01 c	25.19 ± 0.04 c
SPC-5	1.47 ± 0.01 b	2.40 ± 0.03 b	30.56 ± 0.03 b

Sample label explanation: SPC—HUS treatment time 0 min; SPC-0.5—HUS treatment time 0.5 min; SPC-2—HUS treatment time 2 min; SPC-5—HUS treatment time 5 min. All experiments were conducted in triplicate, and results are presented as mean ± standard deviation. Values followed by different letters in columns are significantly different (p < 0.05).

).

The highest content of both free and total sulfhydryl groups was observed in the SPC (Table 1). The HUS treatment caused significant decreases (p < 0.05) in the content of both free and total sulfhydryl groups compared with the SPC for all tested exposure times. The decrease may be attributed to the oxidation of sulfhydryl groups by hydrogen peroxide. Namely, the extreme conditions generated during HUS can induce the production of hydrogen and hydroxyl radicals in the gas phase of the cavities, which can further react and produce hydrogen peroxide [31,32,33]. This cavitation-generated hydrogen peroxide may oxidize susceptible functional groups of proteins, sulfhydryl being one of them, leading to the formation of disulfides, hence reducing the number of available sulfhydryl groups [31,34]. Hu et al. [32] and Zheng et al. [34] reported the same effect of the HUS on the content of sulfhydryl groups in the individual fractions of soy protein, glycinin and conglycinin, and alcohol-denatured SPI, respectively.

The sample subjected to the shortest HUS treatment (0.5 min) exhibited the lowest total sulfhydryl group content. Increasing the HUS exposure time up to 5 min promoted protein molecular rearrangement due to the cavitational high pressure and shear forces [35], likely leading to the exposure of previously buried sulfhydryl groups, and resulting in a significant (p < 0.05) increase in total sulfhydryl content. A similar result was obtained by Zheng et al. [34]. The effect of HUS on the content of free sulfhydryl groups aligns with the trend observed in total sulfhydryl groups. By exposing SPC to ultrasonic treatment, the content of free sulfhydryl groups increased significantly (p < 0.05) up to the exposure time of 5 min. The results in this paper indicate that oxidation of sulfhydryl groups to disulfide bonds occurred quickly, while further HUS exposure caused reorganization of proteins, but it was not enough to destroy all newly formed S-S bonds. The results of SDS-PAGE electrophoresis conducted under both reducing and non-reducing conditions (Figure 1) provide further insight into the findings. Specifically, lanes SPC, reducing and non-reducing, display the SPC profile. In both lines, identical bands corresponding to the glycinin—more precisely, to its acidic (M_w = 37–42 kDa) and basic (M_w = 17–20 kDa) polypeptides—are visible. The presence of the same bands under both reducing and non-reducing conditions confirms that disulfide bonds were already disrupted, suggesting that the tertiary structure of glycinin had been denatured prior to electrophoresis [29]. Moreover, bands originating from the β-conglycinin subunit (71; 67; 50 kDa) [30], which are stabilized by hydrophobic interactions, are also observed in the non-reducing electrophoresis of SPC, highlighting the protein denaturation [36]. During the industrial production of SPC, some steps, including toasting and alcohol extraction, can lead to the protein denaturation [31]. This finding aligns with the content of free and total sulfhydryl groups in the SPC, since protein denaturation can lead to conformational change, and thus, exposure of sulfhydryl groups that are otherwise buried in the hydrophobic pockets [33,37].

New bands are not visible on reducing SDS-PAGE; however, the non-reducing SDS-PAGE confirms formation of the new aggregates of higher molecular weight, which are stacked on top of the stacking gel. This is in accordance with the results obtained for the content of sulfhydryl groups, which suggest sulfhydryl interchange and the formation of new S-S bonds. However, the slight increase in free sulfhydryl content in the samples exposed to HUS probe for 2–5 min suggests that sonication apart from S-S interchange induced rearrangement of the protein and, possibly, some S-S bonds breaking. These findings are in line with the results regarding the effect of HUS on the 7S and 11S soybean proteins reported in the literature [32].

Additionally, significant differences were observed in the solubility patterns of SPC and samples exposed to HUS (Supplementary Figure S1). The solubility [38] of the SPC increased from 10 to approximately 50% after ultrasonic treatment, and the solubility curve has a typical U shape. On the other hand, solubility of the SPC did not change significantly over the entire tested pH range (except pH 11). Low solubility of SPC is in accordance with results obtained from electrophoresis, which indicated denaturation of SPC. Additionally, in the non-reducing conditions, intensity of the band belonging to the basic polypeptide was lower compared with the samples exposed to HUS (lanes SPC-0.5, SPC-2 and SPC-5). The reason might be formation of insoluble precipitates in which basic polypeptides have a prominent role, which is in accordance with the literature data [36]. On the other hand, HUS seems to be an effective way to enable stable dispersion of these basic polypeptides, as visible from the band intensity in lanes 6–8, along with the increased solubility of HUS-treated samples. Denaturation of glycinin and conglycinin can produce smaller fractions that, under the influence of HUS, can form aggregates, which explains the appearance of intense bands of the SPC-0.5, SPC-2 and SPC-5 lines corresponding to molecular masses of approximately 12 kDa. These results align with the literature reports that during ultrasonic treatment, it is possible to achieve interactions between the glycinin and conglycinin subunits, which result in the formation of both soluble and insoluble aggregates [33,37].

Mixing is the first stage in the development of dough for baked goods production. During this stage, changes occur in the formation of inter- and intramolecular disulfide bonds that define the characteristics of the final product. In addition to disulfide bonds, other types of non-covalent interactions, especially hydrophobic interactions, significantly influence and are important for the development of the product structure [39]. In this regard, the effect of ultrasonic exposure time on the hydrophobicity of SPC was investigated, and the results are given in Table 1. SPC exhibits the highest surface hydrophobicity, 34.17 ± 0.02 µgBPB/mg protein. As previously mentioned, harsh conditions during SPC production damage the structures of glycinin and conglycinin, increasing the exposure of hydrophobic residues and enhancing interactions with the dye bromophenol blue [32,40]. Apparently, HUS has significant influence (p < 0.05) on the surface hydrophobicity of soy proteins. Exposure to HUS first decreases, and then, with prolonged exposure, increases surface hydrophobicity. The lowest hydrophobicity is found in the sample treated with HUS for 0.5 min, while further exposure to an HUS probe up to 2 min increases hydrophobicity. An even more significant (p < 0.05) increase in the hydrophobicity of the sample is observed after treatment for 5 min, 30.56 ± 0.03 µgBPB/mg protein. These results verified that exposure to an HUS probe leads to intensive S-S bonding, resulting in aggregate formation, reduction in free sulfhydryl group content and shielding of the hydrophobic surface. The conclusion regarding the increased formation of insoluble aggregates during prolonged exposure of SPC to HUS (>2 min) is supported by findings indicating that these samples exhibit slightly reduced solubility (Supplementary Figure S1) compared with those treated for less than 2 min. The application of HUS treatment on soy protein isolates, particularly at power levels between 0 and 800 W and low-frequency treatment (20 kHz), profoundly enhances their hydrophobicity [41,42]. It can be concluded that HUS treatment may alter proteins’ surface hydrophobicity, making it a powerful method for enhancing protein functionality and performance.

3.2. Influence of HUS Treatment Time on Raman Spectra of Soy Protein Concentrate

The Raman spectrum was analyzed to evaluate the impact of HUS treatment time on the microenvironment, the chemistry of the amino acid side chains within the polypeptide backbone, and the secondary structure [32]. The findings are illustrated in Figure 2, highlighting the Raman spectrum’s key features, and in

Table 2. Normalized intensity values at selected regions of Raman spectra (tyrosyl doublet, phenylalanine band, S-S stretching band and δ-CH band) of SPC exposed to HUS probe (amplitude—30% and frequency—20 ± 0.2 kHz).

Sample Label	Normalized Intensity * of Raman Bands
	Tyr (642.5 cm−1)	Fermi Doublet of Tyrosine (I850/I830)	Phe Band (618.6 cm−1)	Tryptophan Indole-Ring (756 cm−1)	S-S Stretching Band (500–511 cm−1)	CH2 Bending (1439.8–1453.2 cm−1)
SPC	0.148 ± 0.014 a	1.118 ± 0.019 c	0.115 ± 0.006 c	0.234 ± 0.006 c	0.087 ± 0.013 b	1.557 ± 0.054 c
SPC-0.5	0.204 ± 0.016 a	1.346 ± 0.018 a	0.160 ± 0.001 a	0.357 ± 0.017 a	0.165 ± 0.017 a	3.083 ± 0.205 a
SPC-2	0.187 ± 0.017 a	1.271 ± 0.031 a,b	0.135 ± 0.006 b	0.293 ± 0.018 b	0.143 ± 0.018 a,b	2.316 ± 0.216 b
SPC-5	0.173 ± 0.012 a	1.193 ± 0.030 b,c	0.142 ± 0.001 b	0.308 ± 0.013 a,b	0.126 ± 0.013 a,b	2.188 ± 0.131 b,c

* Intensity at the Raman shift (cm⁻¹), shown in parentheses, was normalized to the intensity of the phenylalanine band at 1004 cm⁻¹. Sample label explanation: SPC—HUS treatment time 0 min; SPC-0.5—HUS treatment time 0.5 min; SPC-2—HUS treatment time 2 min; SPC-5—HUS treatment time 5 min. All experiments were conducted in triplicate, and results are presented as mean ± standard deviation. Values followed by different letters in columns are significantly different (p < 0.05).

.

The Tyr doublet ratio (I₈₅₀/I₈₃₀) indicates whether Tyr residues are exposed or buried and highlights the acceptor state of the tyrosine phenolic group; meanwhile, the Raman band intensity at 1439.8–1453.2 cm⁻¹ reflects the aliphatic amino acid side chain microenvironment [27,43]. In the present work, significant (p < 0.05) changes in the normalized intensity of Tyr doublet ratio upon exposure of SPC to HUS were found (Table 2). Namely, the lowest ratio (I₈₅₀/I₈₃₀) was found in the SPC (0.9 < I₈₅₀/I₈₃₀ < 1.45), indicating that the tyrosine was exposed to water/polar environments. The increased I₈₅₀/I₈₃₀ ratio in the samples SPC-0.5 and SPC-2 suggests weakening of hydrogen bonds involving the tyrosine hydroxyl group and consequential conformational changes that allow Tyr residues to adopt more flexible orientations. Additionally, an increase in this ratio can be related to the reorientation of the tyrosine aromatic ring [44,45,46]. The Raman band intensity at 1439.8–1453.2 cm⁻¹ follows a similar trend. The lowest intensity of the bands corresponding to CH₂ bending vibrations was found in the SPC, suggesting the exposure of aliphatic amino acids to the polar environment [47]. However, the exposure of SPC to HUS probe for 0.5 min increased this band intensity, suggesting that the aliphatic side chains became more tightly packed, probably embedded in the protein complex due to the structural rearrangement or aggregate formation. Prolonged HUS treatment of SPC 2–5 min again resulted in exposure of the aliphatic amino acids, suggesting that some aggregates were dissociated during HUS probe treatment, and a decrease in the intensity of the bands corresponding to CH₂ bending vibrations was observed as well as an increase in free sulfhydryl groups content, as reported above [27].

The location of Trp residues according to the band intensity at 756 cm⁻¹ varied among the samples analyzed. The intensity band at 756 cm⁻¹ was found to be lowest in the SPC compared with SPC exposed to HUS. Namely, the highest intensity at 756 cm⁻¹ was found in the sample exposed to HUS probe for 0.5 min. This is a clear indication that the Trp residues tend to become buried inside the protein complex upon 0.5 min sonication [27] in accordance with the intensity of the CH₂ band. These results suggest that exposure to HUS for 0.5 min led to the formation of aggregates and protein rearrangement, with hydrophobic groups being embedded in the protein complex. Supporting this conclusion, a decrease in surface hydrophobicity was observed in the HUS-treated sample. Additionally, according to the results obtained for the sulfhydryl group content, new S-S bonds were formed and probably were responsible for the protein rearrangement. However, prolonged sonication, especially 5 min, induced a decrease in the intensity band near 760 cm⁻¹, meaning that the Trp residues became more exposed. The Trp intensity band trend upon sonication aligns with the hydrophobicity of the SPC samples exposed at different times to the HUS probe. The most hydrophobic was the SPC, and the least hydrophobic sample was the one obtained after exposure to an HUS probe for 0.5 min. Further exposure to the HUS probe, 2 and 5 min, increased the hydrophobicity and decreased the 756 cm⁻¹ band intensity, suggesting Trp exposure towards the hydrophilic environment and structural rearrangement, as already mentioned.

The S-S stretching band positioned in the Raman spectra at 500–511 cm⁻¹ gives insight into disulfide bridges in the soy protein, which are a very important factor in dough stability and development since they are known to differ in stability. The intensity of these bands varied significantly (p < 0.05) among the samples (Table 2). Namely, the lowest band intensity at 500–511 cm⁻¹ was found in the SPC with the highest amount of free sulfhydryl groups. The band intensity strongly increased upon 0.5 min sonication of SPC. The higher exposure to the HUS probe (>0.5 min) resulted in band intensity decrease, suggesting more free sulfhydryl, which aligns with the results above. The distinct frequencies associated with the S-S stretching vibrations in the Raman spectra were categorized into three conformational forms. To quantify these conformational forms in SPC samples, the disulfide band region 490–550 cm⁻¹ was extracted and analyzed by Gaussian function [27,48], and the results are given in

Table 3. Disulfide bond configuration of SPC exposed to HUS probe (amplitude—30% and frequency—20 ± 0.2 kHz).

Sample Label	S-Sg–g–g, %	S-Sg–g–t, %	S-St–g–t, %
SPC	57.25 ± 4.43 a	0.58 ± 0.04 c	42.17 ± 1.74 b
SPC-0.5	34.84 ± 3.46 b	42.69 ± 3.99 b	22.47 ± 2.04 c
SPC-2	21.52 ± 2.05 c	76.07 ± 3.75 a	2.41 ± 0.08 d
SPC-5	11.58 ± 0.95 c	3.62 ± 0.09 c	84.80 ± 2.43 a

Sample label explanation: SPC—HUS treatment time 0 min; SPC-0.5—HUS treatment time 0.5 min; SPC-2—HUS treatment time 2 min; SPC-5—HUS treatment time 5 min. All experiments were conducted in triplicate, and results are presented as mean ± standard deviation. Values followed by different letters in columns are significantly different (p < 0.05).

.

As previously demonstrated, the SPC used in this study was highly denatured, indicating disruption of disulfide bonds. This disruption allowed for the formation of new disulfide bonds when exposed to HUS. Table 3 illustrates that the exposure of SPC to HUS resulted in changes to the conformation of disulfide bonds, which are crucial for dough stability [49]. Based on this observation, it is possible to make some predictions on how the addition of SPC affects the dough’s stability. It is noticeable that SPC exposure to an HUS probe contributed to a decrease in the content of the g–g–g conformation and an increase in the g–g–t and t–g–t conformations, which is consistent with research results available in the literature [40]. Upon exposure of SPC to HUS for 5 min, a significant decrease in the g–g–g conformation was observed, accompanied by an increase in the more flexible g–g–t and t–g–t conformations, indicating increased accessibility of disulfide bonds and favoring intermolecular cross-linking and aggregate formation [50].

The secondary structure of the protein used in dough fortification is fundamental in terms of its impact on starch retrogradation, i.e., on the product’s shelf life and quality [51]. In that regard, the changes in the secondary structure before and upon sonication were analyzed, and the results are given in

Table 4. Secondary structure composition (%) of SPC exposed to HUS probe (amplitude—30% and frequency—20 ± 0.2 kHz).

Sample Label	α-Helix, %	β-Sheet, %	β-Turn, %	Random Coil, %
SPC	12.72 ± 0.07 d	79.83 ± 3.01 a	5.71 ± 0.04 c	1.74 ± 0.04 b
SPC-0.5	33.52 ± 1.58 a	56.80 ± 4.40 b	9.67 ± 0.10 b	n.d.
SPC-2	25.03 ± 1.36 b	63.90 ± 2.89 b	11.07 ± 0.08 b	n.d.
SPC-5	19.71 ± 0.06 c	56.31 ± 3.90 b	16.38 ± 1.23 a	7.60 ± 0.95 a

Sample label explanation: SPC—HUS treatment time 0 min; SPC-0.5—HUS treatment time 0.5 min; SPC-2—HUS treatment time 2 min; SPC-5—HUS treatment time 5 min. All experiments were conducted in triplicate, and results are presented as mean ± standard deviation. Values followed by different letters in columns are significantly different (p < 0.05). Note: n.d. = not determined.

and Figure 3.

Based on the data presented in Table 4, it can be concluded that the α-helix and β-sheet are the predominant secondary structures in SPC, as previously reported in the literature [27]. However, these findings confirm the denatured state of SPC, as indicated by the notably lower α-helix and higher β-sheet contents compared with those of purified glycinin and β-conglycinin [43]. Upon exposure of SPC to the HUS probe, a significant increase (p < 0.05) in α-helix content was observed.

The increase in α-helix content appears to be related to protein solubility in this study and in the study previously reported by Bai et al. [52]. Further exposure of SPC to HUS probe for 2 and 5 min resulted in α-helix content decrease accompanied by a slight rise in β-sheet content corresponding to a reduction in protein solubility. An increase in exposure time (>2 min) decreased the β-sheet content compared with SPC. These findings are consistent with previously reported results concerning the influence of HUS on both native and commercial soy protein isolates [21,34]. Similarly, a decrease in α-helix content and an increase in β-sheet content upon sonication were also observed during the HUS treatment of zein [53].

Compared with SPC, exposure to HUS resulted in an increased content of unordered structures such as β-turns. The highest β-turn content was observed in SPC-5. The increase in β-turn content from unmodified SPC to SPC-5 indicates enhanced molecular flexibility and a transition from ordered to less ordered structural conformations [54]. The trend observed in random coil content followed a pattern similar to that of β-turns. Specifically, exposure of SPC to HUS resulted in a pronounced increase in random coil content after 5 min of sonication (7.60 ± 0.95%). Overall, the findings indicate that HUS induced structural rearrangement and promoted the formation of protein aggregates characterized by a greater proportion of β-structures relative to α-helices [55]. The results of this study suggest that exposure of SPC to HUS led to the structural conversion of β-sheets, β-turns, and random coils into a more ordered, hydrogen bond–stabilized α-helix conformation. Moreover, HUS treatment contributed to the reconstitution of internal hydrogen bonding within the SPC. Thus, ultrasonic processing was demonstrated to be a practical approach for enhancing the structural order of SPC in a manner comparable to that observed in mantle proteins of scallops [56].

3.3. The Influence of HUS-Modified SPC on the Thermo-Mechanical Properties of Enriched Dough

Incorporating proteins into wheat flour changes the dough’s rheological properties, affecting the type of dough and the final product [57]. This study examines the possibility of using the power of HUS technology to modify a single protein for multiple purposes. The Mixolab device and the standard “Chopin+ protocol” investigate how SPC and SPC exposed to HUS influence dough rheology.

Table 5. Mixolab^TM parameters for wheat dough fortified with SPC exposed to HUS probe for different times (amplitude—30% and frequency—20 ± 0.2 kHz).

Sample Label	WA, %	DDT, min	S, min	C2, Nm	C3, Nm	C4, Nm	C5, Nm
Control	52.6 ± 0.85 b	1.98 ± 0.04 b,c	8.7 ± 0.18 a	0.533 ± 0.02 a	2.302 ± 0.05 b	2.247 ± 0.03 a	4.178 ± 0.05 a
SPC	57.6 ± 0.33 a	1.12 ± 0.09 a	8.7 ± 0.30 a	0.525 ± 0.50 a	2.057 ± 0.04 c	1.987 ± 0.22 a,b	3.538 ± 0.54 a,b
SPC-0.5	58.4 ± 0.71 a	0.98 ± 0.01 c	8.9 ± 0.00 a	0.447 ± 0.01 a	1.830 ± 0.14 b	1.759 ± 0.01 c	2.614 ± 0.03 e
SPC-2	58.1 ± 0.71 a	0.95 ± 0.01 c	8.6 ± 0.62 a	0.448 ± 0.02 a	1.860 ± 0.23 b	1.761 ± 0.01 c	2.753 ± 0.03 d,e
SPC-5	57.6 ± 0.42 a	1.32 ± 0.16 b,c	8.2 ± 0.00 a	0.480 ± 0.03 a	1.923 ± 0.19 b	1.828 ± 0.02 c	2.790 ± 0.02 d
	C1-2		C3-2		C3-4		C5-4
Control	0.574 ± 0.04 a		1.769 ± 004 c		0.055 ± 0.00 c		1.931 ± 0.02 a
SPC	0.596 ± 0.02 a		1.532 ± 0.14 a		0.070 ± 0.03 a		1.551 ± 0.13 b
SPC-0.5	0.608 ± 0.05 a		1.383 ± 0.03 d		0.071 ± 0.00 c		0.855 ± 0.04 c
SPC-2	0.620 ± 0.00 a		1.412 ± 0.01 d		0.099 ± 0.00 c		0.992 ± 0.06 c
SPC-5	0.648 ± 0.04 a		1.443 ± 0.06 d		0.095 ± 0.00 c		0.962 ± 0.06 c

Sample label explanation: SPC—HUS treatment time 0 min; SPC-0.5—HUS treatment time 0.5 min; SPC-2—HUS treatment time 2 min; SPC-5—HUS treatment time 5 min. All experiments were conducted in triplicate, and results are presented as mean ± standard deviation. Values followed by different letters in columns are significantly different (p < 0.05).

provides an overview of the key rheological parameters.

The exposure of SPC to HUS led to significant alterations in the protein hydrophobicity, sulfhydryl profile, and secondary structure composition. As a result, SPC altered the rheological properties of enriched dough. Specifically, adding SPC to wheat flour increased the dough’s water absorption capacity, supporting the findings that SPC with low solubility binds more water than initially anticipated [41]. Exposing SPC to HUS for 0.5 and 2 min increased the dough’s water absorption capacity, which can positively affect product yield [58]. Dough fortification with SPC-5 slightly reduced the water absorption capacity; however, these values are still higher than those of the control dough. These results suggest that the increase in soy protein solubility by ultrasonic treatment shows a positive relationship with the water absorption capacity of the dough enriched with HUS-modified SPC. The water absorption capacity significantly impacts (p < 0.05) the dough development time. The control dough had a development time of 1.98 ± 0.04 min, significantly (p < 0.05) longer than the 1.12 ± 0.09 min needed for the dough enriched with SPC to reach maximum consistency. Therefore, it can be concluded that the SPC addition weakened the gluten network, leading to a softer and less elastic dough [59]. Adding SPC-0.5 and SPC-2 reduced the dough development time, probably due to the increment in available water owing to the higher solubility of these SPC types. However, this trend changes with flour fortification using SPC-5. SPC-5 exhibited lower solubility and bound less water than SPC-0.5 and SPC-2, resulting in an increased dough development time.

Different SPC types affect the stability profile of fortified dough in distinct ways. The dough fortified with the SPC-0.5 had the highest stability, while decreases were observed in dough fortified with SPC, SPC-2, and especially, SPC-5. During the first phase of dough development, it is possible to establish various covalent and non-covalent bonds between gluten and soy proteins [60]. Soy protein has some free thiol groups that can interact with gluten proteins, incorporating them into the gluten network. However, it is also possible to establish hydrophobic interactions between hydrophobic gluten and soy protein. Both interactions rule the stability of the dough [49]. Dough fortified with the SPC had stability similar to the control flour (Table 5). This SPC had the higher content of the most stable gauche–gauche–gauche (g–g–g) disulfide bond conformation [49]. The dominance of this bond in the protein structure may reduce the likelihood of bond cleavage during the dough processing, and therefore, potentially contribute to the mechanical stability of the dough matrix. Although the highest content of free sulfhydryl groups was shown to be present in this SPC, it is assumed that the driving force for interactions with gluten proteins is hydrophobicity, as the highest hydrophobicity among the analyzed samples was exhibited by this SPC. These results are in line with previous results, where it was established that intermolecular g–g–g confirmation is decisive for dough stability, while the hydrophobic interactions are important in the structural evolution of dough [49].

The lowest stability was observed in doughs enriched with SPC-2 and SPC-5, and this can be associated with significantly (p < 0.05) changed proportions of disulfide bond conformations. Namely, these samples contain more disulfide bonds that are energetically less stable, g–g–t and t–g–t, and hence, are more prone to strain during the dough processing [49]. In this case, where the hydrophobicity of SPC-2 and SPC-5 is lower than that of SPC, a different mechanism for the establishment of bonds between soy proteins and gluten can be expected. Although the content of free sulfhydryl groups is still lower than in SPC, the higher proportion of unstable disulfide bond conformations means that more frequent breakage can be expected and that an increase in the proportion of free sulfhydryl groups may be observed. Cross-linking of these sulfhydryl groups with gluten sulfhydryl groups leads to dilution of the gluten network and a decrease in dough stability [61].

Among all the samples analyzed, the best stability was demonstrated by the dough enriched with SPC-0.5. The functional properties were analyzed, and it was shown that the highest solubility was exhibited by this sample compared with all other samples. It is suggested that less aggregation was experienced and steric hindrance for gluten hydration was reduced. As a result, enough available water was retained by the gluten to allow the proper development of a stable network. Further, Raman analysis of the proteins’ disulfide bond conformations revealed that much more stable disulfide bond structures were present in the proteins prepared in this way, indicating resistance to mixing and lower free sulfhydryl content available for cross-linking with gluten, and therefore, a lower possibility of interfering with regular gluten network development. Additionally, it was demonstrated that the exposure of the hydroxyl group of tyrosine was significantly (p < 0.05) increased by HUS treatment in this case (I₈₅₀/I₈₃₀ = 1.346 ± 0.018), thereby enhancing the probability of intermolecular hydrogen bond formation between SPC and gluten [62,63]. In the case of the dough enriched with SPC-0.5, a key role in the stabilization of the dough is likely played by the exposure of tyrosine residues and the formation of hydrogen bonds, despite the low hydrophobicity and the limited number of free sulfhydryl groups. The importance of hydrophobic interactions and hydrogen bonds in the formation of the gluten network has also been observed in the zein–gluten system [64].

SPC addition to wheat flour significantly changed protein strength and resistance during heating (C1-2). The speed of protein weakening (C1-2) in the dough enriched with SPC increased with HUS treatment time, following the order SPC-0.5 > SPC-2 > SPC-5, indicating that prolonged sonication enhances protein softening. This increase in C1-2 can be the consequence of gluten network dilution, where protein molecules are packed less compactly and the speed of protein weakening during mixing and heating is higher [65]. The doughs enriched with the SPC are more resistant to weakening caused by heat and mixing. This can be related also to the specificity and the type of the bonds (S-S bonds, hydrophobic interactions, types of disulfide bond conformation, hydrogen bonding) formed between gluten molecules and the SPC [66].

By observing the data in Table 5, we conclude that the addition of SPC and ultrasonically modified SPC significantly (p < 0.05) affects the starch gelatinization rate (C3-2). The starch gelatinization rate in the control was the highest among all analyzed samples. SPC is in a highly dehydrated state and can bind more water than usual; therefore, the competition between starch for available water is favored. As a result, the starch granule swelling is retarded, and the system’s viscosity decreases. In contrast, SPC exposed to HUS has different effects on the starch gelatinization rate. The gelatinization rate significantly increased (p < 0.05) after enrichment of the dough with SPC-0.5, SPC-2, and SPC-5. SPC-0.5 showed the highest solubility; therefore, the highest competition rate between SPC-0.5 and starch granule exists, there is a reduced amount of water for starch to swell, and the gelatinization rate decreases. The slight decrease in solubility for SPC-2 and SPC-5 increased slightly the gelatinization rate.

The amylase activity (C3-4) in the control and the doughs fortified with the SPC and HUS-treated SPC follows the trend of the starch gelatinization rate. Amylase activity in the control dough yielded dough with viscosity 0.055 ± 0.00 Nm. The addition of SPC in the dough changed the dough viscosity by increasing the ability of starch to swell and, subsequently, by increasing the amylase activity. Adding foreign protein to the dough sometimes acts as a physical barrier, blocking the amylase active center and inhibiting its activity, thus reducing the dough’s viscosity [51]. In this case, the presence of HUS-modified SPC increased amylase activity through a second mechanism. Namely, during this phase, dough temperature increased, inducing soy protein denaturation and release of water, which became available for starch to swell [67].

Furthermore, the addition of SPC-0.5 positively influenced the starch gelatinization rate and the amylase activity. In this dough, the amylase activity was increased compared with the control dough, and this increasing trend remained unchanged in the sample enriched with SPC-2, wherefrom the amylase activity declined after the addition SPC-5, since the addition of these samples reduced the rate of starch gelling. This effect is related to the lower solubility of this SPC sample compared with SPC-0.5 and SPC-2.

Finally, the starch retrogradation rate of the dough was remarkably changed after dough supplementation with SPC, especially with HUS-treated SPC, as evidenced by C5-4 (Table 5). The highest starch retrogradation rate (C5-4) was observed in the control dough. This was reduced by the addition of SPC to the wheat flour. Adding protein to wheat flour can either speed up or slow down the process of starch retrogradation. The impact of this process is directly related to the specific groups present on the surface of the protein and the interactions with starch. The slowing of starch retrogradation in dough enriched with SPC can be directly attributed to its highly pronounced hydrophobic character. The hydrophobic nature of the protein leads to the repulsion of water molecules from the starch granules, which plays a crucial role in releasing and reassociating amylose [51]. Moreover, the amylose double helix has a hydrophobic center, which is susceptible to interacting with hydrophobic compounds such as SPC and forms stable complexes and prevents amylose associations [51]. Incorporating SPC-0.5, SPC-2, and SPC-5 into flour significantly slows the retrogradation process and effectively extends the product’s shelf life. The mechanism behind this process can be the explained by the solubility of soy proteins exposed to ultrasonic treatment for 0.5, 2, and 5 min. Namely, under the above conditions, an increased solubility of the mentioned proteins was recorded, notably expanding the dough’s water-binding capacity. Also, the presence of proteins with a high water-binding capacity positively affects water mobility during starch retrogradation; more precisely, it prevents water mobility and syneresis. In this way, recrystallization is prevented. The second factor affecting the starch retrogradation is the composition of the secondary structure. More specifically, a higher content of β-secondary structure was observed in these samples, formed at the expense of α-helices, which is usually accompanied by the formation of a densely packed protein network that can also prevent water access to starch and recrystallization [51]. Furthermore, helices are the most stable secondary structure, since hydrogen bonding between chains provides the rigidity of molecular structure, and therefore, forms stable and rigid structures between protein and starch molecules, preventing amylose chains from aligning and impeding retrogradation [68]. As shown, the SPC-0.5 has the highest content of α-helixes, which are, along with β-turns, rigid; both form similar bonds with starch and give the same effect on the amylose recrystallization. In contrast, β-sheets and random coils are more flexible structures, and their effects on starch retrogradation depend on the specific protein–starch interactions that are established [69]. Figure 4 shows the Mixolab Profiler output, which is provided as a complementary visualization of the results summarized in Table 5.

4. Conclusions

This study demonstrates that HUS is an effective tool for modifying the SPC, whose functionality and structure have been significantly altered by production conditions. Control of the treatment time is crucial for inducing desired modifications, including changes in free sulfhydryl group content, accessibility of hydrophobic regions, secondary structure composition, and disulfide bond conformation. The changes in SPC solubility were reflected in the dough’s water absorption capacity and development time. Interactions between SPC and gluten protein were driven by protein hydrophobicity and the type of disulfide bond conformation, which, in turn, influenced the dough stability. Moreover, the ratio of ordered and disordered protein structures influenced the starch retrogradation, thereby influencing textural changes during storage and contributing to the retention of product freshness. Overall, HUS enables targeted modification of SPC, allowing a single protein ingredient to be tailored for application in a wide range of functional flour-based products, highlighting the commercial potential of the methodology given in this work. Further research could focus on applying HUS to diverse protein sources and assessing the sensory and nutritional properties of the resulting bakery products.