Effects of Ultrasonication Combined with Enzymatic Treatment on the Structure and Function of Soy Protein Isolate

Abstract

Natural soy protein isolate (SPI) exhibits suboptimal functional characteristics, including limited solubility, reduced foaming capacity, and diminished emulsifying ability. Conventional singular-modification techniques are unable to enhance multiple functional properties concurrently, thereby posing challenges in fulfilling the varied requirements of food processing. Therefore, this study employed ultrasonic and pepsin enzymatic modification techniques on SPI. By varying ultrasonic frequency (20 kHz, 207 kHz) and sonic energy density (295 W/L, 590 W/L), different modified protein samples were obtained. The effects of single treatment, combined treatment, and varying ultrasonic parameters on their structure and functionality were investigated. The results indicate that compared to single enzymatic hydrolysis, combination-treated SPI exhibited reduced fluorescence intensity and UV absorbance, along with significant decreases in methionine (Met) and free-sulfhydryl (SH) content (p < 0.05). Particle size decreased while distribution became more uniform, and relative molecular weight also diminished. This indicates that combined processing induces more pronounced changes in the protein’s primary to higher-order structures, thereby enhancing functional properties. Specifically, surface hydrophobicity (H₀) and emulsification stability (ESI) improved, while emulsifying capacity (EAI) significantly increased (p < 0.05). In summary, ultrasonication combined with enzymatic hydrolysis exhibits synergistic effects, optimizing protein structure and functional characteristics. This approach facilitates the development of functional foods and broadens their application scope.

1. Introduction

Proteins constitute fundamental components of human cells and tissues, performing essential physiological functions such as regulating metabolism, supporting immune defense, and maintaining muscle strength, making them crucial for human health. However, with population growth and improved living standards, protein resource shortages have become increasingly severe [1]. Compared to animal proteins, plant proteins not only align better with healthy dietary trends but also offer sustainability advantages [2]. Among these, soy protein isolate (SPI) is extensively employed because of its superior processing characteristics, rich nutrients, and cost-effectiveness, making it one of the primary sources for replacing animal protein [3]. SPI encompasses all essential amino acids and exhibits digestibility comparable to animal protein [4]. The structural characteristics of SPI determine its functional properties. β-conglycinin (7S) and glycinin (11S), which account for approximately 65–80% of the overall protein content, make up the majority of the protein composition [5]. These proteins adopt a rigid and compact globular conformation under the influence of covalent and non-covalent interactions [6]. This results in SPI exhibiting poor molecular flexibility, low solubility, weak emulsifying properties, and limited foaming capacity, thereby restricting its applications in food production [7]. Therefore, employing appropriate methods to alter the SPI structure and improve its functional characteristics is of paramount importance.

Currently, physical methods (like heat treatment and high-pressure processing), chemical methods (like pH adjustment and oxidation) [8], and biological methods (such as enzymatic and fermentation approaches) comprise the majority of traditional SPI modification techniques. However, each of these methods has inherent limitations: Physical modification yields limited effects, primarily involving non-covalent interactions. Structural changes are reversible, and functional improvements are often insignificant. Additionally, treatments involving high temperatures or intense irradiation may degrade flavor or destroy protein nutrients [9,10]. Chemical methods pose safety risks and environmental-pollution concerns due to chemical reagent usage [11]. Biological methods suffer from limited applicability, stringent reaction conditions, and poor product stability. The application of composite modification technology addresses the shortcomings of individual methods by leveraging synergistic effects, representing a highly effective strategy.

In recent years, ultrasonic technology has attracted growing interest as an environmentally friendly, efficient, and non-thermal physical approach [12]. Ultrasound, a mechanical wave with frequencies exceeding the human auditory threshold (>20 kHz), primarily modifies proteins through cavitation effects. During ultrasonication, cavitation bubbles form within liquid systems. These bubbles undergo continuous expansion and contraction over time, culminating in a violent collapse that generates localized extreme conditions: temperatures (5000 K), pressures (100 MPa), and high shear forces [13,14]. The combined action of these factors disrupts non-covalent connections, including hydrogen bonds and hydrophobic interactions, which causes protein subunits to dissociate and aggregate to achieve modification [15]. Additionally, the cavitation effect triggers water molecule dissociation and secondary reactions, producing extremely reactive free radicals (·OH, H·, etc.) that interact with protein molecules [5]. To improve the effectiveness of ultrasonic treatment, several studies have focused on combining ultrasonication with other modification methods. Liurong Huang et al. treated SPI with dual-frequency ultrasound and ionic liquids. Results indicated that combined treatment affected SPI’s non-covalent interactions, thereby altering its emulsifying properties [16]. Nevertheless, studies combining ultrasonication and enzymatic hydrolysis for SPI modification remain scarce, with most research focusing on enzymatic conditions. Therefore, investigating the combined ultrasonic–enzymatic treatment and its ultrasonic parameters may offer potential benefits for enhancing SPI’s physicochemical properties [17,18].

Enzymatic hydrolysis is a method where proteases cleave specific chemical bonds in substrates under optimal pH and temperature conditions, breaking down macromolecular proteins into smaller ones [19]. It has attracted considerable interest owing to its specificity, environmental sustainability, high efficiency, and capacity to effectively enhance functional properties [20]. Research indicates that factors such as degree of hydrolysis and enzyme type significantly influence protein modification. Restriction enzyme digestion does not fully hydrolyze proteins into small peptides or amino acids; instead, it cleaves specific peptide bonds at designated sites within the protein, generating amphiphilic protein peptides [21]. The effects of hydrolysis duration (1, 2, 3, and 4 h) and enzyme type (pepsin, bromelain, and trypsin) on SPI structure and function were examined by Yiting Gao et al. Modified hydrolysates (SPHs) were blended with guar gum (GA) as emulsifiers for emulsion preparation. Experimental results revealed that SPH/GA emulsifiers exhibited reduced particle sizes, increased negative charges, elevated levels of interfacially adsorbed proteins, and enhanced stability [22]. Compared to neutral and alkaline proteases, pepsin is an acidic protease that specifically breaks down peptide bonds between aromatic amino acids in protein molecules. Its enzymatic products contain a higher proportion of hydrophobic peptide segments, and it is widely available and inexpensive. However, pepsin has limited hydrolysis sites, making it difficult to achieve deep hydrolysis with a single enzyme. Additionally, the hydrophobic peptide segments tend to aggregate, hindering improvements in functional properties. Combining pepsin with ultrasonication may potentially resolve these issues.

Therefore, this work aims to explore how ultrasonication and enzymatic hydrolysis affect the structural and functional properties of SPI. First, pepsin was used to hydrolyze SPI, followed by enzyme inactivation. The hydrolysate was then subjected to ultrasonic treatment under different ultrasonic frequencies (20 kHz, 207 kHz) and sound energy densities (295 W/L, 590 W/L). Subsequently, structural alterations were analyzed by measuring endogenous fluorescence spectra, Fourier transform infrared (FTIR) spectra, amino acid composition, particle size distribution, and SDS-PAGE. Functional effects of different modification methods were then evaluated through surface hydrophobicity (H₀), emulsifying capacity (EAI), and emulsion stability (ESI) studies. Finally, correlation analysis established relationships between structural modifications, surface hydrophobicity, emulsifying capacity, and emulsion stability of modified SPI. This approach overcomes limitations of single-modification methods and, by investigating ultrasonic parameter effects on protein structure and function, offers novel insights for green, efficient regulation of soybean protein functional properties.

2. Materials and Methods

2.1. Materials

Soy protein isolate (SPI, protein content ≥ 90%) was purchased from Solarbio Technology Co., Ltd. (Beijing, China). Pepsin (enzyme activity ≥ 1200 U/g) was supplied by Sinopharm Chemical Reagent Co., Ltd. (Shanghai, China). Emulsifying oil was sourced from COFCO Donghai Grain & Oil Industry Co., Ltd. (Suzhou, China). Unless otherwise specified, all additional chemical reagents employed in this investigation were of analytical grade purity.

2.2. Sample Preparation

Powdered SPI was dissolved in water at a solid-to-liquid ratio of 10:1 (mg/mL) and was stirred magnetically at room temperature for 2 h. The solution pH was then adjusted to 2.0 with 2 M HCl. Pepsin (300 U/g SPI) was subsequently added to the solution and mixed thoroughly. The solution was incubated at 37 °C for 30 min. Following incubation, pH was modified to 7.0 using 2 M NaOH, and the solution was heated to 90 °C for 10 min to stop the enzymatic activity. The sample was freeze-dried for subsequent experiments [23]. In order to explore the influence of combined ultrasonic and enzymatic treatment and ultrasonic parameters on SPI, the above freeze-dried powder was dissolved in deionized water (10 mg/mL) and then treated under different ultrasonic frequency and acoustic energy density conditions. Low-frequency ultrasonication (20 kHz) was conducted utilizing an ultrasonic cell disruptor fitted with a 10 mm diameter probe (XO-1000D, Nanjing ATPIO Co., Ltd., Nanjing, China). For each treatment, 90 mL and 45 mL of the enzymatic hydrolysate were processed, respectively. The probe was immersed 1 cm below the liquid surface and sonicated for 40 min at acoustic power densities of 295 W/L and 590 W/L, using intermittent pulse mode (5 s on, 5 s off). To avoid temperature rise and protein denaturation during ultrasonication, the beakers were submerged in an ice-water bath. For mid-frequency ultrasonication (207 kHz), an ultrasonic generator equipped with a double-walled beaker (amplitude 5 Vpp, Nantong Longyi Electronic Technology Co., Ltd., Nantong, China) was used. To achieve acoustic power densities of 295 W/L and 590 W/L, 50 mL and 25 mL of the enzymatic digestion solution were placed in the double-walled beaker, respectively. Ultrasonication was performed for 20 min using a continuous-pulse mode. A circulating water bath maintained a constant temperature within the jacketed beakers during ultrasonication. The combined ultrasonicated and enzymatically treated sample solutions were freeze-dried for subsequent use. Control protein samples were prepared using the identical protocol, excluding the steps of enzymatic digestion and ultrasonication.

2.3. Intrinsic Fluorescence Spectroscopy

A fluorescence spectrophotometer (LS55, PerkinElmer, Waltham, MA, USA) was utilized to measure the sample solution’s endogenous fluorescence spectrum. The solution was diluted with deionized water to a level of 0.067 mg/mL for analysis. The excitation wavelength was set at 280 nm, the emission wavelength scanning range was set at 300–500 nm, and the slit width was set at 5 nm [24].

2.4. UV-Vis Spectrometry

The method described by Shizhang Yan et al. was slightly modified. The sample solution was adjusted with deionized water to achieve a level of 0.2 mg/mL. A dual-beam UV-vis spectrophotometer was utilized to detect the UV spectrum. Deionized water was first utilized as a blank control for baseline correction, followed by scanning the test solution in the 200–720 nm wavelength range [3].

2.5. Fourier Transform Infrared Spectroscopy (FTIR)

Freeze-dried protein samples were ground with KBr powder at a 1:100 (w/w) ratio, then pressed into thin pellets using a tablet press. The Fourier transform infrared (FTIR) spectra of the samples were acquired utilizing a Nicolet IS50 FTIR spectrometer (Thermo Fisher Scientific, Dreieich, Germany) at 25 °C. The wavenumber range was 400–4000 cm⁻¹, with 16 scans at a resolution of 4 cm⁻¹. Data were processed and analyzed utilizing PeakFit software (version 4.12, SYSTAT Software Inc., San Jose, CA, USA) and OriginPro 2021 (version 9.8, OriginLab Corporation, Northampton, MA, USA), revealing patterns of secondary structure changes in proteins [25].

2.6. Amino Acid Composition Analysis

Types and concentrations of amino acids in the sample solution were analyzed utilizing an ultra-high-pressure liquid chromatograph (Infinity 1290, Agilent Technologies, Santa Clara, CA, USA). First, the sample was treated with acid hydrolysis to hydrolyze proteins into amino acids. A 10 mL vacuum hydrolysis tube was filled with a 3 mL aliquot of the sample solution (10 mg/mL) and 3 mL of 6 M HCl. Excess gas was purged using a nitrogen purge device until the liquid surface reached a gentle boil; then the sample was hydrolyzed at 110 °C for 20 h. A 0.22 μm aqueous membrane was used to filter the hydrolysate after it had been collected and diluted five times with deionized water. Amino acids in the sample were detected using Agilent standard protocols and automatic OPA derivatization [26]. Chromatographic analysis was conducted utilizing an Agilent Poroshell C-18 column with dimensions of 4.6 × 100 mm. After being combined in a certain ratio, mobile phases A and B were eluted at 1.5 mL/min. Mobile phase A (2 L, pH 8.2) contained 2.8 g disodium hydrogen phosphate (Na₂HPO₄) and 7.6 g sodium borate decahydrate (Na₂B₄O₇·10H₂O). Mobile phase B (1 L) was a mixture of acetonitrile (chromatographic-grade), methanol (chromatographic-grade), and deionized water in a 45:45:10 volume ratio. The mobile phases were filtered through a 0.45 μm aqueous membrane (MCE, 47 mm diameter, Tianjin Jinteng, Tianjin, China) and a 0.45 μm organic membrane (Nylon, 47 mm diameter, Tianjin Jinteng, China) using a vacuum filtration apparatus (1 L receiver flask, 300 mL funnel, Tianjin Jinteng, China), respectively, followed by ultrasonication for 15 min to remove bubbles from the mobile phase. The gradient elution procedure was configured as follows: 0 to 0.35 min, 2% solvent B; 0.35 to 13.4 min, 57% solvent B; 13.4 to 13.5 min, 100% solvent B; 13.5 to 15.7 min, 100% solvent B; 15.7 to 15.8 min, 2% solvent B; and 15.8 to 18 min, 2% solvent B. The flow rate was set at 1.5 mL/min, and the injection volume was set at 1 μL. Excitation and emission wavelengths for the fluorescence detector were set at 340 and 450 nm, respectively. Subsequently, peak areas corresponding to each amino acid were applied to their respective standard curves to quantify the concentration of each amino acid within the sample [27].

2.7. Particle Size Distribution

To compare particle size distribution among different treatment groups, measurements were performed utilizing a nanoparticle size analyzer (ZS90, Malvern Analytical Ltd., Malvern, UK). Sample solutions were diluted to a level of 1 mg/mL with deionized water, thoroughly mixed, and then injected into the sample cell. After equilibration at 25 °C for 60 s, particle size (Z-average size) and distribution curve data for the protein samples were measured and exported. Parameter settings were as follows: protein refractive index 1.45, protein absorbance 0.001, water refractive index 1.33, water viscosity 0.8872 cP. All tests were repeated three times [28].

2.8. SDS-PAGE

SDS-PAGE can reveal the relative molecular weight distribution of proteins and changes in intermolecular interactions (such as C-C and S-S bonds). Conventional Tris-SDS-PAGE electrophoresis can only resolve large macromolecular proteins, exhibiting extremely low resolution for proteins with smaller molecular weights, particularly those below 10 kDa. In contrast, Tricine-SDS-PAGE effectively separates proteins and peptides with molecular weights ranging from 1 to 10 kDa, making it the primary method for denaturing separation of peptides via electrophoresis.

First, the protein samples were denatured by mixing the sample solution (10 mg/mL) with loading buffer at 1:1 (v/v). The mixture was vortexed and then heated in boiling water for 5 min to fully unfold the protein structure. After cooling, the mixture was centrifuged, and the resulting supernatant was used for spot application. Since untreated proteins have relatively high molecular weights, Tris-SDS-PAGE electrophoresis is employed. A total of 5 μL of protein supernatant was added to the lane of a 12.5% resolving gel. Electrophoresis was performed at 110 V for 2 h. After electrophoresis, the gel was stained with Coomassie Brilliant Blue G-250 staining solution for 1 h. Destaining was completed with deionized water, after which the gel was photographed and analyzed [29]. For modified histones with relatively low molecular weights, Tris-SDS-PAGE yielded poor separation, necessitating Tricine-SDS-PAGE. The electrophoresis chamber (16.5% separating gel, 10% interlayer gel, 4% concentrating gel) was placed in an ice-water bath. Anode buffer was added to the outer chamber, and cathode buffer was added to the inner chamber. Pre-electrophoresis was performed at 30 V for 10 min. Then, 5 μL of protein supernatant was added to the loading wells. Electrophoresis was carried out at 30 V until the leading edge of the band reached the top of the separating gel. Subsequently, the voltage was increased to 100 V, and electrophoresis was continued until the band was 1–2 cm above the bottom of the gel. Electrophoresis was then stopped, and subsequent staining, destaining, and photographing were performed [30].

2.9. Free-Sulfhydryl (SH) Content

The DTNB assay was utilized to measure the amount of free SH groups. This technique involves the reaction of DTNB with free SH groups in the sample, producing a compound with specific light absorption characteristics at a defined wavelength. The absorbance measured at this wavelength is then used to calculate the free-sulfhydryl content via a standard curve. The protein solution was diluted to a level of 1 mg/mL with deionized water. Then, 1 mL of the diluted solution was taken and was thoroughly mixed with 4 mL of Gly-Tris buffer (0.086 M Tris, 0.09 M Gly, 4 mM Na₂EDTA, pH 8.0). Subsequently, 50 μL of Ellman’s reagent (4 mg/mL, DTNB dissolved in Gly-Tris buffer) was added, and the mixture was thoroughly mixed and allowed to react at room temperature in the dark for 1 h. A mixture without protein samples was used as a blank control. The absorbance was measured at 412 nm utilizing a UV spectrophotometer (UV-1700, Shimadzu, Kyoto, Japan), and free-SH content was calculated according to the following formula [31].

SH (μmol/g) = 10⁶ × A × D/(1.36 × 10⁴ × b × C)

where A denotes the absorbance at 412 nm, D denotes the dilution factor, C denotes the sample concentration (1 mg/mL), b denotes the path length (1 cm), 1.36 × 10⁴ denotes the molar extinction coefficient (L·mol⁻¹·cm⁻¹), and 10⁶ denotes the conversion factor from mol to μmol.

2.10. Surface Hydrophobicity (H₀)

8-Anilino-1-naphthalenesulfonic acid (ANS) was utilized as a fluorescent probe to measure the surface hydrophobicity of protein samples. A range of concentrations of protein solutions were made, including 0.02, 0.04, 0.06, 0.08, and 0.1 mg/mL. Subsequently, 4 mL of protein solution was thoroughly combined with 20 μL of ANS solution (8 mM) and incubated in the dark for 20 min. Following reaction completion, fluorescence intensity was detected utilizing a fluorescence spectrophotometer (LS55, PerkinElmer, Waltham, MA, USA) at an excitation wavelength of 390 nm and an emission wavelength of 470 nm. Finally, a relationship curve was plotted with protein concentration and fluorescence intensity. The curve slope represents the surface hydrophobicity [32].

2.11. Emulsifying Properties

In total, 4 mL of protein solution (2 mg/mL) was mixed with 1 mL of oil. The mixture was homogenized using a homogenizer (T18, IKA, Guangzhou, China) at 10,000 rpm for 2 min. Then, 50 μL of emulsion was collected from the sample’s bottom at 0 min and 10 min, and it was immediately added to 5 mL of SDS solution (0.1%, w/v). After thorough mixing, the absorbance was measured at 500 nm utilizing a UV spectrophotometer (UV-1700, Shimadzu, Kyoto, Japan) [33]. The formulas below were utilized to figure out the emulsifying activity index (EAI) and emulsion stability index (ESI).

EAI (m²/g) = 2 × 2.303 × A₀ × DF/[C × ∅ × (1 − θ) × 10,000]

ESI (min) = A₀ × 10/(A₀ − A₁₀)

where C is the protein solution concentration (g/mL), θ is the volume fraction of oil in the emulsion (0.2), DF is the dilution factor, ∅ is the optical path length (1 cm), 10,000 is the conversion factor from cm² to m², 10 is the time interval (min), and A₀ and A₁₀ are the emulsion’s absorbance values at 0 and 10 min, respectively.

2.12. Statistical Analysis

All experiments were repeated three times, and results are expressed as mean ± standard deviation. Differences between data sets were assessed for significance utilizing one-way analysis of variance (ANOVA) with SPSS 26.0 software (IBM, Armonk, NY, USA). Statistical significance was determined using a p-value of less than 0.05. All figures and tables were generated using OriginPro 2021 software.

3. Results and Discussion

3.1. Impact of Combined Ultrasonic and Enzymatic Treatment on the Structure of SPI

3.1.1. Analysis of the Tertiary Structure of SPI with Different Modifications

Protein tertiary-structure changes can be reflected via endogenous fluorescence spectroscopy. This is because the intrinsic fluorescence of chromophores is extremely sensitive to variations in their microenvironment’s polarity [34]. When the maximum absorption wavelength (λ_max) exceeds 330 nm, the microenvironment surrounding Tryptophan (Trp) is polar, with longer λ_max values indicating stronger microenvironmental polarity [35]. Fluorescence intensity changes in SPI across different treatment groups are shown in Figure 1A. Trp residues are exposed to polar environments since all samples exhibit maximum absorption wavelengths exceeding 330 nm. The untreated SPI sample exhibited a maximum absorption wavelength of approximately 355 nm. In contrast, both enzymatic digestion alone and the combined treatment of SPI resulted in red-shifted maximum absorption wavelengths, shifting from 355 nm to 375 nm and 365 nm, respectively. Furthermore, compared to the untreated group, the fluorescence intensity of SPI modified by different methods was significantly enhanced, suggesting that the protein’s tertiary structure underwent unfolding, exposing the Trp residues and hydrophobic groups originally buried within the protein molecule [36]. During SPI enzymatic hydrolysis, pepsin acts on hydrophobic amino acids, including Tryptophan (Trp), Tyrosine (Tyr), and Phenylalanine (Phe), increasing their exposure and thereby enhancing their fluorescence intensity [23]. During ultrasonic treatment, the cavitation effect generated by ultrasound alters the protein’s tertiary structure, exposing hydrophobic regions within the molecule [37]. After combined treatment, the fluorescence intensity of SPI shows a downward trend with the decrease in ultrasonic frequency, which is the opposite of the change in acoustic energy density. This may be because the increased exposure of hydrophobic groups will promote protein aggregation to form a more stable structure [31].

Figure 1.

Endogenous fluorescence spectra (A) and UV–visible spectra (B) of SPI from untreated, enzymatic treatment, and composite treatment groups (20 kHz-295 W/L, 20 kHz-590 W/L, 207 kHz-295 W/L, 207 kHz-590 W/L).

Changes in protein tertiary structure, such as the microenvironment around aromatic amino acids and the relative dynamics of side-chain amino acid residues, can also be described using ultraviolet–visible spectroscopy [3]. Figure 1B, consistent with endogenous fluorescence spectroscopy results, shows the absorbance of modified SPI within the ultraviolet absorption bands of Phe, Tyr, and Trp (240–320 nm). The modified SPI exhibited significantly increased absorbance relative to the untreated control group. Furthermore, the absorbance of the composite-treated group decreased with decreasing ultrasonic frequency and increasing sonic energy density. The increase in absorbance may stem from exposure of hydrophobic regions or alterations in its secondary structure resulting from ultrasound treatment. However, excessive exposure of hydrophobic domains can give rise to protein reaggregation, which in turn manifests as a decrease in absorbance [5,38].

3.1.2. Analysis of the Secondary Structure of SPI with Different Modifications

Chemical bonds and secondary structural components of proteins were examined using Fourier Transform Infrared Spectroscopy (FT-IR). Figure 2A presents FT-IR spectra corresponding to various protein samples. SPI exhibits four characteristic peaks: 3280.85 cm⁻¹ (amide A), 1627.52 cm⁻¹ (amide I), 1519.10 cm⁻¹ (amide II), and 1228.19 cm⁻¹ (amide III) [39]. Among these, the most prominent amide I vibration band is a sensitive region for protein secondary structure [40], with α-helix, β-sheet, β-turn, and random-coil regions located at 1650–1660 cm⁻¹, 1600–1640 cm⁻¹, 1660–1700 cm⁻¹, and 1640–1650 cm⁻¹, respectively [29]. The characteristic peaks in the modified group resembled those in the untreated group, indicating that the modification did not cause the protein to create new chemical bonds. However, the amide absorption peak intensity in SPI was enhanced following both single and combined treatments, indicating alterations in the protein’s secondary structure and the exposure of hydrophobic groups from the inside to the surface of the protein molecules [41]. To further clarify variations in protein secondary-structure content, deconvolution analysis of the amide I band yielded calculated percentages for α-helices, β-sheets, β-turns, and random coils (Figure 2B). β-sheets and β-turns constitute the predominant secondary structures in native SPI, while α-helix content is minimal. Compared to the single-treatment group, combined medium-frequency ultrasonication and enzymatic modification led to a decrease in α-helix and β-turn content of SPI, accompanied by an increase in β-sheet content. Their reduction indicates a transition of the protein molecule from an ordered, compact state to a disordered, relaxed state [42]. This may occur because ultrasonic treatment disrupts intermolecular interactions within the protein, while the resulting cavitation effect unfolds the SPI structure, promoting the conversion of α-helices into β-folds and random coils, thereby enhancing the disorder of the protein structure [43]. This aligns with findings by Lianzhou Jiang et al., who observed decreased α-helix and increased β-sheet proportions in black bean protein isolate (BBPI) following ultrasonication [44]. Conversely, the low-frequency-ultrasonication combined with enzymatic-digestion group exhibited opposite patterns, likely due to more intense cavitation at lower frequencies causing protein depolymerization and rearrangement, thereby altering secondary structure content. Furthermore, regardless of whether ultrasonication at 20 kHz or 207 kHz was combined with enzymatic digestion, higher sonic energy density resulted in more disordered protein structures. This indicates that high sonic energy density more significantly disrupts intermolecular interactions within proteins.

Figure 2.

FT-IR (A) and secondary structure distribution (B) from untreated, enzymatic treatment, and composite treatments (20 kHz-295 W/L, 20 kHz-590 W/L, 207 kHz-295 W/L, 207 kHz-590 W/L). Significant differences (p < 0.05) are marked by distinct letters within columns.

3.1.3. Analysis of the Primary Structure of SPI with Different Modifications

Protein primary structure is composed of amino acids, encompassing the types, quantities, sequence orders, and linkage patterns of amino acid residues. Changes in amino acids reflect the exposure levels of hydrophobic amino acids in different sample groups and ultrasonically induced intramolecular redox reactions. Figure 3A and Figure 3B present the HPLC chromatograms and amino acid content bar charts for SPI from different treatment groups, respectively. Fifteen amino acids were detected in untreated SPI: aspartic acid (Asp), glutamic acid (Glu), serine (Ser), histidine (His), glycine (Gly), threonine (Thr), arginine (Arg), alanine (Ala), tyrosine (Tyr), valine (Val), methionine (Met), phenylalanine (Phe), isoleucine (Ile), leucine (Leu), and lysine (Lys). Among these, Glu was the most abundant, while Met was the least abundant, corroborating the results reported by Chamba et al. [45]. Compared to untreated samples, all amino acid contents in SPI decreased significantly after modification (p < 0.05). Notably, Met content in the composite treatment group also showed a significant reduction compared to the single treatment group (p < 0.05). The effect of medium-frequency combined enzymatic hydrolysis was more pronounced. Instead, peak areas between Thr (Peak 6) and Arg (Peak 7) significantly increased. This behavior could be attributed to oxidation products of Met (e.g., methyl mercaptan or methyl mercaptansulfonate) [46]. Ultrasonication induces cavitation in liquids, generating instantaneous localized extreme temperatures, pressures, and shear forces. While these conditions can unfold protein structures, they may also degrade released free amino acids or small peptides, particularly those sensitive to oxidation, heat, or mechanical stress. Furthermore, cavitation generates highly reactive free radicals such as hydroxyl radicals (·OH) and hydrogen radicals (H·), leading to oxidative degradation of Met. Yue Wu et al. (2021) further investigated the oxidative rate of Met over time at a frequency of 355 kHz and an intensity of 2.14 W/mL, demonstrating the dominant role of chemical effects during the initial phase of medium-frequency ultrasonic treatment [27].

Figure 3.

HPLC chromatograms (A) and corresponding amino acid content profiles (B) of untreated, enzymatically treated, and composite-treated (20 kHz-295 W/L, 20 kHz-590 W/L, 207 kHz-295 W/L, 207 kHz-590 W/L) SPI. (A: 1 = Asp, 2 = Glu, 3 = Ser, 4 = His, 5 = Gly, 6 = Thr, 7 = Arg, 8 = Ala, 9 = Tyr, 10 = Val, 11 = Met, 12 = Phe, 13 = Ile, 14 = Leu, 15 = Lys. B: Distinct lowercase letters denote statistical differences among treatments, with a significance threshold of p < 0.05.

3.1.4. Particle Size Distribution of SPI in Different Treatment Groups

The degree of protein aggregation is reflected in particle size, which is an essential metric for assessing protein stability. A higher propensity for protein aggregation is indicated by larger particle sizes, which therefore impacts functional characteristics [47]. Figure 4A and Figure 4B illustrate the mean particle size and the particle size distribution of SPI under untreated, single-treatment, and combined-treatment conditions, respectively. As shown in Figure 4A, the average particle size of SPI significantly decreased after modification compared to the untreated sample (p < 0.05), reducing from 232.83 nm to 179.13 nm, 100.55 nm, 91.34 nm, 175.57 nm, and 174.63 nm, respectively. Notably, the composite-modified SPI exhibited smaller average particle sizes than those from single enzymatic modification, with the 20 kHz low-frequency ultrasound treatment yielding more pronounced effects (p < 0.05). These results may relate to alterations in intermolecular interactions such as hydrogen bonds, hydrophobic forces, and electrostatic interactions. Enzymatic processing or cavitation effects from ultrasonication disrupt the forces between protein aggregates, effectively reducing particle size [23]. Furthermore, the physical effects of low-frequency ultrasound (including shear forces and microjet effects) are more intense than those of medium-frequency ultrasound [48], resulting in the smallest SPI particle size when low-frequency ultrasound was combined with enzymatic treatment. As shown in Figure 4B, the untreated and single-treated groups exhibit broad peak distributions, indicating poor SPI particle uniformity. In contrast, the combined treatment yields a narrower particle size distribution, demonstrating that ultrasound enhances the uniformity of enzymatically processed SPI particles. Additionally, the peak position shifts to the left, confirming that combined treatment results in smaller SPI particle sizes. Mokhtar Dabbour et al. reported similar findings, observing reduced particle sizes in sunflower protein after limited enzymatic hydrolysis and ultrasonic modification [49].

Figure 4.

Particle size distribution (A) and distribution profile (B) of SPI subjected to untreated, enzymatic treatment, and composite treatment (20 kHz-295 W/L, 20 kHz-590 W/L, 207 kHz-295 W/L, 207 kHz-590 W/L). Electrophoresis pattern of untreated SPI (C). Electrophoresis patterns of SPI subjected to enzymatic treatment and composite treatment (20 kHz-295 W/L, 20 kHz-590 W/L, 207 kHz-295 W/L, 207 kHz-590 W/L) (D). (A: Distinct lowercase letters denote statistically significant differences among the various treatments (p < 0.05). D: Non-Reducing: 1: Enzymatic; 2: 20 kHz-295 W/L; 3: 20 kHz-590 W/L; 4: 207 kHz-295 W/L 5: 207 kHz-590 W/L. Reducing: 1′: Enzymatic; 2′: 20 kHz-295 W/L; 3′: 20 kHz-590 W/L; 4′: 207 kHz-295 W/L 5′: 207 kHz-590 W/L.)

3.1.5. Molecular-Weight (MW) Distribution of SPI in Different Treatment Groups

SDS-PAGE is primarily utilized to ascertain relative MW and to reflect the degree of protein degradation or aggregation. Figure 4C shows electrophoresis patterns of native SPI under non-reducing and reducing conditions. Untreated SPI exhibits a molecular-weight distribution primarily between 10 and 180 kDa, with distinct bands for 7S and 11S clearly observable. The 7S fraction is characterized as a trimeric glycoprotein consisting of α′, α, and β subunits, which exhibit approximate MWs of 71 kDa, 67 kDa, and 50 kDa, respectively [50]. 11S is a hexameric glycoprotein comprising acidic subunit A and basic subunit B with MWs of approximately 35 kDa and 20 kDa, respectively [51]. Non-reducing and reducing electrophoresis results for different modification treatment groups are shown in Figure 4D. Single-modified and composite-modified samples exhibited similar band patterns. Following modification, SPI demonstrated significantly enhanced band intensity in the low-molecular-weight region, primarily concentrated below 30 kDa. Concurrently, the band for the basic subunit B (~20 kDa) became noticeably thinner and weaker within this range, clearly demonstrating that the combined action of enzymatic digestion and ultrasonication degraded SPI subunits into fragments with smaller relative molecular weights. This phenomenon arises because the modification process disrupts covalent or non-covalent connections between protein molecules, contributing to protein degradation. This result is in line with what Xixi Wu et al. found [23].

3.1.6. Changes in the Free Sulfhydryl Groups (SH) of SPI with Different Modifications

Disulfide bonds represent essential covalent linkages that contribute significantly to the structural stability of proteins. The disruption of these bonds can result in protein denaturation, thereby revealing previously buried sulfhydryl groups within the protein interior [52]. Therefore, free-sulfhydryl content serves as a key indicator for evaluating protein conformational changes and functional properties [53]. Figure 5A illustrates the variation in free-sulfhydryl content across different treatment groups. Compared to untreated SPI, all modified samples exhibited a significant rise in free-sulfhydryl content (p < 0.05). This is because the compact globular structure of native SPI encloses its sulfhydryl groups internally, resulting in low free-sulfhydryl content. However, following modification, the enzyme digestion and ultrasonic cavitation effects disrupt the protein’s compact globular structure, causing structural unfolding and disulfide bond breakage. This exposes the internally buried sulfhydryl groups to the molecular surface [54,55]. However, except for the 20 kHz-590 W/L treatment group, the combined-treatment groups exhibited a significantly smaller increase in free-sulfhydryl content compared to the single-treatment groups (p < 0.05). This phenomenon may stem from hydrogen peroxide (H₂O₂) generated during ultrasonication, which facilitates the oxidation of exposed free sulfhydryl groups, thereby decreasing their content [56]. Alternatively, ultrasonication may cause protein aggregation, re-encapsulating some free sulfhydryl groups [57]. Notably, variations in acoustic energy density during ultrasonication significantly impacted free-sulfhydryl content, which decreased markedly with reduced energy density (p < 0.05). This phenomenon can be ascribed to the diminished cavitation effect at low acoustic power densities, but it may be more likely to promote oxidative reactions. Ultrasound-generated free radicals (such as ·OH) are more readily reactive with sulfhydryl groups at lower energies, oxidizing them into disulfide bonds or other oxidative products.

Figure 5.

Free-sulfhydryl content (A), surface hydrophobicity (B), and emulsifying activity and stability (C) of SPI subjected to untreated, enzymatic treatment, and composite treatments (20 kHz-295 W/L, 20 kHz-590 W/L, 207 kHz-295 W/L, 207 kHz-590 W/L). Distinct lowercase letters represent significant differences between different treatments (p < 0.05).

3.2. Impact of Combined Ultrasonic and Enzymatic Treatment on the Function of SPI

3.2.1. Changes in Surface Hydrophobicity (H₀) of SPI with Different Modifications

Protein tertiary structural changes are reflected in surface hydrophobicity (H₀), which is closely linked to the extent of hydrophobic group exposure. These changes influence functional properties such as protein conformation, solubility, and gelling capacity [58]. Changes in H₀ between native SPI and modified SPI are shown in Figure 5B. Untreated SPI exhibits low H₀, attributed to its native globular structure, where hydrophobic groups are buried within the protein molecule. Compared to the untreated group, all modified samples exhibited significantly enhanced H₀ (p < 0.05), increasing from 67.37 ± 0.72 to 138.48 ± 12.17, 140.65 ± 5.32, 136.50 ± 4.09, 141.77 ± 8.98, and 144.79 ± 4.29, respectively. This indicates that the modification treatment induced irreversible conformational changes in SPI, exposing its internal hydrophobic groups [59]. This phenomenon was further corroborated by findings from endogenous fluorescence spectroscopy (Figure 1A) and UV–visible spectroscopy (Figure 1B). Additionally, except for the 20 kHz-590 W/L treatment group, the composite-treatment groups exhibited a slight improvement in H₀ compared to the single-treatment groups. This result may be attributed to the cavitation effect induced by ultrasound, which further disrupts the spherical conformation of proteins, thereby exposing more hydrophobic groups [60]. However, under conditions combining enzymatic hydrolysis with 20 kHz-590 W/L ultrasonication, the surface hydrophobicity of the samples slightly decreased. This may be because the intense ultrasonication conditions enhanced intermolecular interactions among proteins, leading to protein repolymerization and the re-encapsulation of some hydrophobic groups within the molecules. Shizhang Yan et al. also noted that as ultrasonic power increases, proteins can reform aggregates, resulting in reduced H₀ [3].

3.2.2. Changes in the Emulsifying Properties of SPI with Different Modifications

Proteins’ emulsifying qualities are analyzed utilizing the emulsifying activity index (EAI) and emulsifying stability index (ESI). EAI reflects a protein’s capacity to form water–oil interfaces, while ESI reflects its capacity to maintain emulsion stability and resist shear stress [54]. Figure 5C illustrates the changes in emulsifying characteristics of SPI after being unmodified, single-modified, and combination-modified. Ultrasonication combined with enzymatic hydrolysis significantly increased the EAI value of SPI compared to untreated and single-treated samples (p < 0.05), with more pronounced effects observed at lower sonic energy densities. The sample treated with enzymatic hydrolysis followed by 20 kHz-295 W/L ultrasonication exhibited the highest EAI value (3.14 m²/g). Additionally, the modification slightly increased SPI’s ESI value, with the highest ESI value (16.01 min) observed in samples treated with enzyme digestion combined with 207 kHz-590 W/L ultrasonication. This suggests that exposed hydrophobic groups were absorbed at the interface, generating a more stable bilayer, even if its EAI value is lower than that of other samples [61]. Figure 6 also clearly shows that the oil–water interface disappears after homogenization, forming an emulsion, and the modified emulsion exhibits greater uniformity and stability. The enhanced emulsifying performance can be attributed to protein unfolding during ultrasonication. Changes in secondary structure manifested as reduced α-helix content and increased β-sheet content, simultaneously improving protein flexibility. This promotes interactions between proteins and lipid molecules while altering protein arrangement at the oil–water interface [38]. Research by Ying Zhu et al. also indicates that increased protein flexibility enhances its capacity to stabilize the water–oil interface [62]. According to other studies, greater flexibility raises surface hydrophobicity and solubility. Improved surface hydrophobicity promotes interactions between protein molecules within the interfacial layer, constructing a dense interfacial layer that consequently influences emulsification properties, while enhanced solubility facilitates protein distribution at the interface [3]. However, when the sound energy density reached 590 W/L, the emulsifying activity of SPI showed a declining trend. This indicates that high sound energy density may result in the exposure of numerous hydrophobic groups, which could facilitate protein aggregation via hydrophobic interactions [63].

Figure 6.

Photographs of untreated, enzymatically treated, and composite-treated (20 kHz-295 W/L, 20 kHz-590 W/L, 207 kHz-295 W/L, 207 kHz-590 W/L) SPI before (A) and after (B) emulsification.

3.3. Correlation Analysis

A correlation analysis of physicochemical properties was performed on various protein samples to evaluate the connection between protein structure and function (see Figure 7). In the figure, values ranging from −1 to 1 represent the Pearson correlation coefficient (r). Positive correlation is shown by the red area, while negative correlation is shown by the green area. The intensity of color corresponds to the level of correlation. First, there was a significant negative association between EAI and particle size (p ≤ 0.01) and a significant positive association between ESI and free-SH concentration (p ≤ 0.05). Increased free-SH content results from disulfide bond cleavage. Protein flexibility is increased by higher free-SH content, which facilitates protein rearrangement at the interface and increases links between proteins and lipids [64]. Conversely, reduced particle size indicates a larger specific surface area of droplets, which enhances EAI [65]. Second, H₀ showed a significant negative correlation with particle size, in contrast to free-SH content (p ≤ 0.01). Studies indicate that during modification, intermolecular interactions within proteins are disrupted, causing tertiary-structure unfolding. This exposes internal hydrophobic groups and sulfhydryl residues while reducing particle size. Additionally, particle size distribution exhibited a significant negative correlation with α-helix and β-turn content (p ≤ 0.05). This might be explained by the disruption of intermolecular hydrogen bonds and hydrophobic interactions within SPI molecules under shear forces generated by enzymatic digestion and ultrasonic cavitation. This process leads to the depolymerization of macromolecular aggregates into smaller peptide fragments, directly resulting in reduced protein particle size. Meanwhile, α-helices and β-turns—as regular secondary structures within SPI molecules maintained by hydrogen bonds—underwent rearrangement of hydrogen bonds during aggregate depolymerization and peptide chain cleavage. This transformation of the originally loose, disordered coiled structure ultimately manifested as increased α-helix and β-turn content.

Figure 7.

Correlation between structure and function properties of SPI with different modifications (A,B). * represents significant differences between different indicators (*: p ≤ 0.05, **: p ≤ 0.01, ***: p ≤ 0.001, ****: p ≤ 0.0001).

3.4. Mechanism of Modification Treatment for SPI

Figure 8 illustrates the mechanism of SPI modification. Restriction enzyme digestion is a modification technique that selectively cleaves specific peptide bonds in soy protein under mild conditions using proteases such as alkaline protease, trypsin, or flavor protease. Unlike conventional complete enzymatic hydrolysis, this method partially hydrolyzes proteins into functional peptides by precisely controlling enzyme type, enzyme–substrate ratio, and hydrolysis time. Its key advantage lies in exposing protein active sites to enhance solubility (solubility in the isoelectric point range increases by 80%) and emulsifying properties, while preventing the accumulation of bitter peptides and loss of gelatinous texture caused by excessive hydrolysis. This technology enables the targeted production of active peptides with specific molecular weights (3–20 kDa), offering dual advantages: enhanced functional properties (e.g., doubling foaming capacity) while preserving nutritional value. Combining it with physical modification techniques (such as ultrasonic-assisted enzymatic hydrolysis) further boosts efficiency [23].

Figure 8.

Proposed mechanisms of enzymatic and ultrasound-induced SPI modification. ((A): Enzymatic and ultrasonic treatments enhance the emulsibility of SPI. (B): Pepsin selectively cleaves peptide bonds in SPI, breaking large proteins into smaller peptides with varying molecular weights, thereby exposing active sites and improving emulsification. (C): Low-frequency ultrasound induces cavitation, where bubble implosion generates extreme local high temperature, pressure, and shear forces, disrupting intermolecular interactions and unfolding protein structure to expose hydrophobic groups. Medium-frequency ultrasound primarily acts via sonochemical effects, where bubble collapse dissociates water into hydroxyl radicals and hydrogen radicals. Hydroxyl radicals react with aromatic amino acids, causing oxidation that potentially alters surface charge distribution and hydrophobicity.)

The cavitation effect is the core mechanism of ultrasonic processing technology, essentially involving the dynamic evolution of microscopic bubbles generated in liquid media under ultrasonic influence. When ultrasonic waves propagate through a liquid, they create localized low-pressure zones during the negative-pressure phase. Upon exceeding the liquid’s cavitation threshold, the cohesive forces between liquid molecules break down, producing micrometer-scale cavitation bubbles. These bubbles rapidly collapse and violently implode during the subsequent positive-pressure phase, with the entire process occurring within nanoseconds. The instantaneous collapse of cavitation bubbles generates extreme physical conditions in the local space, including instantaneous high temperatures (approximately 5000 K), high pressures (approximately 50 MPa), and microjet velocities reaching up to 400 km/h. This extreme environment subjects protein molecules to multiple physical actions: powerful shear forces dissociate protein aggregates, shock waves disrupt secondary intermolecular bonds, and high temperatures promote protein conformational unfolding. Notably, cavitation exhibits significant spatial and temporal inhomogeneity, with its intensity closely related to ultrasonic frequency, power density, and liquid properties. Within the common frequency range of 20–100 kHz, low-frequency ultrasound tends to generate more intense transient cavitation, while high-frequency ultrasound predominantly produces milder steady-state cavitation. This unique energy transfer mechanism makes ultrasonic processing a highly promising technique for physical modification of proteins [66].

Sonochemistry refers to the chemical reaction mechanism generated by cavitation effects during ultrasonic processing. When cavitation bubbles violently collapse under ultrasonic influence, the gas and vapor within undergo dissociation under conditions of extremely elevated temperature and pressure, generating highly reactive free radicals. In aqueous solutions, water molecules thermally decompose within cavitation bubbles to produce hydroxyl radicals (·OH) and hydrogen radicals (H·). These reactive species diffuse into the bulk solution and interact with protein molecules. Hydroxyl radicals possess strong oxidizing properties, capable of attacking specific amino acid residues in proteins—such as aromatic amino acids like Trp, Tyr, and His—leading to oxidative modifications. This oxidation may alter the surface charge distribution and hydrophobicity of proteins while also promoting the cleavage and rearrangement of intramolecular disulfide bonds. During high-power ultrasonication, dissolved oxygen within cavitation bubbles is activated to generate reactive oxygen species like singlet oxygen (¹O₂), further intensifying oxidative reactions. While excessive oxidation may diminish protein nutritional value, moderate sonochemical treatment has the potential to improve functional attributes, including solubility and emulsifying capacity [48].

To overcome the limitations of single-modification approaches, the synergistic effect of ultrasonic and enzymatic modification demonstrates unique advantages. This physical–biological coupling strategy not only maintains high emulsion stability across a pH range of 3–9 but also reduces interfacial adsorption energy by 42%, offering a novel approach for developing environmentally responsive emulsion systems [67].

4. Conclusions

This study demonstrates that compared to after single enzymatic hydrolysis treatment, both the fluorescence intensity and UV absorbance of SPI decreased to varying degrees following combined treatment. These decreases were further amplified as the ultrasonic frequency decreased and the acoustic power density increased. Simultaneously, it induced the rearrangement of protein secondary structures. The simultaneous application of medium-frequency ultrasound and enzymatic hydrolysis resulted in a reduction in α-helix and β-turn structures within SPI, accompanied by an increase in β-sheet content. Conversely, low-frequency ultrasound produced opposite results, and an escalation in sonic energy density corresponded with a greater degree of protein structural disorder. Further investigation revealed that the co-treated group’s Met content significantly declined (p < 0.05), indicating that extreme conditions generated by cavitation effects and highly reactive free radicals during ultrasonication may degrade and oxidize Met, with mid-frequency ultrasound exhibiting a more pronounced effect. These results collectively demonstrate that ultrasonic cavitation alters protein structures from primary to higher levels. Along with a more uniform particle size distribution and a lower relative molecular weight, the combined change also decreased the average particle size of SPI, with a more pronounced effect at 20 kHz (p < 0.05). The conformational changes in the protein further optimized its functional properties. The free-SH content in the combined treatment group decreased significantly with decreasing sonic energy density (p < 0.05), while H₀ and ESI improved, and EAI was significantly enhanced (p < 0.05). The EAI value of the sample treated under 20 kHz-295 W/L ultrasonic conditions reached 3.14 m²/g. This study leveraged the synergistic effects of ultrasonication and enzymatic hydrolysis to overcome the limitations of single-modification methods, achieving more effective improvements in SPI’s structure and function. Examining the mechanisms by which ultrasonic treatment affects protein structure and functional properties offers novel perspectives for the environmentally friendly and efficient modulation of soybean protein functionalities.

Abbreviations

The following abbreviations are used in this manuscript:

SPI	Soy protein isolate
FTIR	Fourier Transform Infrared Spectroscopy
SH	Free sulfhydryl
H0	Surface hydrophobicity
EAI	Emulsifying activity index
ESI	Emulsion stability index
MW	Molecular weight

Author Contributions

Conceptualization, W.G. and Y.W.; methodology, W.G. and Y.M.; software, Y.X.; validation, Y.M. and Y.X.; formal analysis, W.G. and Y.M.; investigation, W.G.; resources, Y.W. and Z.C.; data curation, W.G. and Y.M.; writing—original draft preparation, W.G.; writing—review and editing, Y.M., Y.X. and Y.W.; visualization, W.G.; supervision, Y.W. and Z.C.; project administration, Z.C.; funding acquisition, Y.W. All authors have read and agreed to the published version of the manuscript.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The data presented in this study are available on request from the corresponding authors. The data are not publicly available due to privacy or ethical restrictions.

Conflicts of Interest

The authors declare no conflicts of interest.

Funding Statement

This research was funded by Yue Wu. The APC was funded by Yue Wu. This work was supported by the Youth Fund of the National Natural Science Foundation of China [Grant No. 32402291]; Jiangsu Provincial Youth Fund [Grant No. BK20241575]; China Postdoctoral Science Foundation [Grant No. 2024M761436].