Development and characterization of high internal phase emulsions for lycopene encapsulation with safflower seed meal globulin modified via ultrasound, heating and pH-shifting
Ke'er Xiao, Lingyu Gao, Qiuyu Lu, Qiaoyu Wang, Ziteng Zhao, Mukaddas Sai, Xinyu Meng, Lili Guan, Jing Yang, Linna Du
TL;DR
This study shows how modifying safflower proteins with ultrasound, heat, and pH changes improves their ability to stabilize emulsions for lycopene delivery.
Contribution
A novel triple-modification strategy (ultrasound, heating, and pH-shifting) enhances safflower seed meal globulin functionality for bioactive delivery.
Findings
UHA-SMG formed stable HIPEs with 75% internal phase volume and good rheological properties.
LYC encapsulation achieved 96.39% efficiency with improved stability against UV, heat, and storage.
Modification increased α-helical content and altered protein surface properties.
Abstract
This study investigated the synergistic effect of ultrasound in combination with pH-shifting and heating modification on the physicochemical properties, structural characteristics, and emulsifying performance of safflower seed meal globulins (SMG). Furthermore, the efficacy of the modified protein (UHA-SMG) as a natural emulsifier was evaluated for preparing lycopene (LYC)-loaded high internal phase emulsions (HIPEs). Under the suitable modification conditions (ultrasonic power: 500 W, ultrasonic time: 5 min, temperature: 70 °C, pH: 9.0), UHA-SMG exhibited improved physicochemical properties, including altered micromorphology, reduced particle size and interfacial tension, as well as enhanced zeta-potential, surface hydrophobicity, wettability, and solubility. Structural characterization via far-UV circular dichroism, UV–Vis spectroscopy, and fluorescence spectroscopy indicated that the…
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TopicsProteins in Food Systems · Microencapsulation and Drying Processes · Pickering emulsions and particle stabilization
Introduction
1
Nowadays, emulsions, thermodynamically unstable suspensions in which one immiscible liquid is dispersed as fine droplets throughout another liquid, are widely recognized as important systems owing to their multiple functionalities [1]. Recent advances in structural design have enabled the engineering of diverse emulsion systems with enhanced and broadened functional properties [2]. Among these, high internal phase emulsions (HIPEs) are promising emulsion systems that contain a dispersed phase volume fraction exceeding 74% [3]. Owing to their distinctive microstructure and physicochemical properties, HIPEs have been extensively utilized in various applications, including drug delivery systems, templates for porous materials, and filter membranes [4]. HIPEs provide a substantial volume fraction of the oil phase, facilitating the incorporation of hydrophobic bioactive compounds. Additionally, their gel-like consistency and enhanced microbiological stability, attributed to low water activity, render them highly suitable for food-related uses. For instance, in a study conducted by Liu et al., HIPEs were prepared to be used as mayonnaise replacers [5]. Therefore, HIPEs present valuable applications in diverse industries, from spreadable food and drug delivery systems to cosmetic emulsions.
Conventional stabilization of HIPEs relies on small-molecule surfactants, which are capable of diffusing in the aqueous medium and adsorbing at oil–water interfaces, thereby preventing drop aggregation and conferring colloidal stability [6]. While widely used, small-molecule surfactants such as Tween 20 and cetyltrimethylammonium bromide possess low-molecular-weight characteristics that lead to several inherent drawbacks. These include reversible adsorption, limited interfacial strength, and susceptibility to Ostwald ripening—factors that compromise emulsion stability, hinder precise control over droplet size and distribution, and ultimately limit long-term storage and practical utility [7]. Further driving the search for alternatives are growing concerns regarding the potential adverse effects of conventional surfactants on human health and the environment. Consequently, significant interest has emerged in developing next-generation stabilizers capable of forming emulsions with superior stability, tunable droplet characteristics, and enhanced safety.
Recently, as essential biomacromolecular polymers, proteins have received extensive attention. These substances, which range from nano- to micro-particles in size, are amphiphilic compounds with excellent interfacial activity [8]. Proteins facilitate emulsion formation by adsorbing onto new oil droplet surfaces, reducing interfacial tension to promote droplet breakup, and ultimately stabilizing the system via steric and electrostatic forces [9]. Thus, proteins are considered as promising emulsifiers for stabilizing emulsion formulations. Compared to animal proteins, plant-based proteins offer superior extractability, cost-effectiveness, greater availability, and enhanced accessibility, thus making them a more sustainable alternative for emulsion stabilization [10]. However, with the exception of a few extensively utilized plant proteins—such as those derived from soybean, pea, mung bean, and chickpea—many other abundant botanical protein sources remain under exploited.
Safflower (Carthamus tinctorius L.), an Asteraceae family member, is a traditional oilseed crop primarily cultivated for its medicinal and edible benefits [11]. It is worth noting that many unsaturated fatty acids (linoleic acid, oleic acid, etc.), and multiple bioactive substances are found in safflower seed oil [12]. Therefore, safflower seeds are considered a promising source of raw materials for high-quality oil production. However, while safflower oil is extensively utilized, the considerable seed meal generated as a by-product represents an under-valued resource that is frequently disregarded [13]. Moreover, it is reported that various active components, such as proteins, fibers, minerals, and other compounds, were found in safflower seed meal [14]. Among these, protein accounts for a significant proportion of safflower seed meal. Yildirim et al. determined that the crude protein content of safflower seed meal to be approximately 20% [15]. In our previous work, globulins (SMG) were prepared from safflower seed meal, and their structural and functional characteristics were evaluated [16]. However, the functionality of plant proteins is often limited by their lower solubility and poorer emulsifying performance relative to commonly employed animal proteins [17]. As is well known, multiple factors might influence the performance of plant proteins, such as composition, molecular size, structure, temperature, pH, etc. For instance, some proteins exhibit a compact molecular architecture wherein numerous functional groups remain buried, thereby limiting their surface activity, solubility, as well as emulsification and foaming capacities [17], [18]. Therefore, enhancing the performance of SMG is essential for their use in developing SMG stabilized HIPEs.
Several techniques have been developed to modify the protein structure to further improve their functional properties [17], [18], [19]. As a green processing method that is easy to operate and low-cost, ultrasonic treatment induces thermal, mechanical, and chemical effects, which can lead to significant alterations in protein structures, thereby altering their functional properties [20], [21]. For instance, the size, surface hydrophobicity of pea protein were improved after ultrasonic treatment [22]. Additionally, other methods are also used for the modification of some proteins, including heat treatment, pH-shifting, enzymatic hydrolysis, etc [19]. Therefore, developing modification methods suitable for globulins derived from safflower seed meal is particularly important for their utilization and development.
This study aimed to enhance the functional properties of SMG through a combined modification approach involving ultrasonication, heat treatment, and pH-shifting. The structural and physicochemical changes of SMG following these modifications were systematically investigated. Subsequently, the HIPEs were fabricated using either native SMG or modified SMG (referred to as UHA-SMG) as stabilizer. A comprehensive comparison was conducted between the two types of emulsions, including their microscopic and visual appearance, microstructure, interfacial properties, rheological behaviors and stability. Furthermore, HIPEs stabilized by UHA-SMG were employed as a delivery system for lycopene (LYC), and their stability was assessed. This work not only provides a foundational basis for broadening the application prospects of safflower seed meal, but also offers essential experimental evidence supporting the utilization of UHA-SMG stabilized HIPEs in food systems.
Materials and methods
2
Materials
2.1
Commercial safflower seed meal (Xinjiang Lincheng Hengda Trading Co., Ltd. China) was finely pulverized using a high-speed crusher (YB-4500A, YUNBANG, China) and then passed through a 60-mesh sieve. Subsequently, defatting of the safflower seed meal was carried out twice with n-hexane (solid–liquid ratio of 1:3 g/mL) at room temperature for 1.5 h, followed by removal of the residual solvent via rotary evaporation.
Preparation of globulins from safflower seed meal
2.2
Safflower seed meal globulins (SMG) were extracted following the method of Xiao et al [16]. Briefly, to eliminate the influence of albumins, the defatted meal was first mixed with distilled water at a ratio of 1:20 (g/mL) and stirred at 1000 rpm for 100 min at 25 °C. After centrifugation at 8000 rpm for 25 min, the resulting precipitate was collected. Subsequently, the precipitate was resuspended in 1.24 mol/L NaCl at a 1:47 (g/mL) ratio. After being stirred at 1000 rpm (37 °C, 110 min), the mixture was centrifuged at 8000 rpm for 15 min. Following collection, the supernatant was subsequently frozen at −80 °C for 24 h, lyophilized, and the resulting powder was sealed and stored at −20 °C.
Selecting the suitable modification parameters for safflower seed meal globulins
2.3
The SMG suspension (1.0% w/v) was formulated by dispersing SMG in deionized water under continuous magnetic stirring at 1000 rpm for 4 h. Then, SMG suspension was modified with ultrasound, heating coupled with pH-shifting, and the influences of modified parameters on the particle size, zeta-potential and solubility of SMG were also investigated. Firstly, the SMG was treated with ultrasound using an ultrasonic bath system (KQ-250DB, KunShan Ultrasonic Technical Co., Ltd., Suzhou, China) at various ultrasonic power (0, 200, 300, 400, and 500 W), and the other parameters were fixed (ultrasonic time 5 min, ultrasonic temperature 25 °C, the pH of suspension 7). After treatment, the particle size, zeta-potential and solubility of SMG were determined. Subsequently, the suitable ultrasonic duration of SMG was selected in the ranges of 5–20 min at constant ultrasonic power (500 W), temperature (25 °C) and pH (7). Similarly, the influence of different temperatures (25, 50, 70, 90, and 100 °C) on the particle size, zeta-potential and solubility of SMG was tested (ultrasonic power 500 W, ultrasonic time 5 min, pH 7). For pH selection, SMG suspensions were ultrasonicated (500 W, 5 min, 70 °C), cooled, and then adjusted to pH 3, 5, 7, 9, and 11. The resulting samples were characterized in terms of particle size, zeta-potential, and solubility. For all the above tests, the ultrasonic frequency was fixed at 40 KHz.
Determining the particle size, zeta-potential and solubility of native and modified SMG
2.4
The particle size and zeta-potential of diluted samples were measured using a Zetasizer Nano (Malvern Instruments Ltd., UK). The solubility of modified and unmodified SMG was measured referring to other studies [23]. Briefly, following centrifugation of the dispersion (8000 rpm, 10 min, 4 °C), the protein concentration in the supernatant was quantified using a BCA assay kit (Solarbio, Beijing, China).
Characterization of native and modified SMG
2.5
Modification process of SMG
2.5.1
Following hydration at 4 °C for 1 h, the SMG suspension (1.0%, w/v) was subjected to ultrasonication treatment at 500 W and 70 °C for 5 min. After cooling to room temperature, its pH was adjusted to 9.0 ± 0.2 using 0.1 mol/L NaOH.
Scanning electron microscopy (SEM) observation
2.5.2
In line with previous studies, the microstructures of native SMG and UHA-SMG were examined using an scanning electron microscope (SU8100, Hitachi High-Technologies, Japan) [24]. The magnification was set at 5000 × and 15000 × .
Ultraviolet visible (UV-Vis) spectrum analysis
2.5.3
The SMG and UHA-SMG were dispersed in phosphate buffered saline (PBS, 0.01 mol/L, pH 8) at a concentration of 0.1 mg/mL. After thorough mixing, a UV–vis spectrophotometer (Evolution 220, Thermo Electron, USA) was employed to scan the absorption spectra of the two samples in the wavelength range of 200–400 nm [25].
Far-UV circular dichroism (CD) spectra
2.5.4
Far-UV CD spectra of both native SMG and UHA-SMG samples were acquired according to the procedure outlined by Peng et al [26]. Shortly, the samples were prepared by dilution to 10 mg/mL in phosphate buffer (0.01 mol/L, pH 7.0) followed by centrifugation (1000 rpm, 15 min). The resulting supernatant was further diluted to 0.1 mg/mL and analyzed using a ChirascanTM spectrometer (Applied Photophysics Ltd., UK). Spectra were recorded at 25 °C across 190–260 nm with instrumental parameters adopted from the literature [26].
Fluorescence spectroscopy
2.5.5
Fluorescence spectra were acquired using an F-4600 spectrophotometer (Hitachi, Japan) on samples (0.1 mg/mL in 0.01 M phosphate buffer, pH 8.0) as described above. Measurements were taken with an excitation wavelength of 290 nm and an emission range of 290–400 nm, a slit width of 2.5 nm, an acceleration voltage to 700 V, and a response time of 0.5 s.
Surface hydrophobicity analysis
2.5.6
To quantify the difference in surface hydrophobicity, the fluorescent probe 8-anilino-1-naphthalenesulfonic acid (ANS) was used to analyze both native and UHA-modified SMG suspensions [27]. Fluorescence intensity (FI) was recorded on an F-7000 fluorescence spectrophotometer (Hitachi, Japan) at room temperature with an excitation wavelength of 390 nm and an emission wavelength of 400-600 nm, a slit width of 10 nm. Surface hydrophobicity (H_0_) was defined as the slope value of the FI against protein concentration (mg/mL).
Interfacial tension (IFT) detection
2.5.7
For the measurement, protein dispersions (0.1% w/v in deionized water) were prepared. The dynamic interfacial tension at the oil–water interface was determined using the pendant drop technique on a DSA30S tensiometer (KRÜSS, Germany) [28]. Briefly, a pendant droplet (≈45 μL) of the protein solution was formed in a quartz cuvette filled with rice bran oil. During the 3600 s measurement, droplet images were continuously captured and digitally processed. The interfacial tension was calculated by fitting the contour to the Young-Laplace equation.
Formation and characterization of UHA-SMG-stabilized high internal phase emulsions (HIPEs)
2.6
Preparation of HIPEs
2.6.1
HIPEs were prepared by homogenizing corn oil, which was purchased from the local market, and an aqueous phase containing either native SMG or UHA-SMG using a homogenizer (T18BS25, IKA, Germany) at 15,000 rpm for 2 min. To determine the optimal oil volume fraction, HIPEs stabilized by native SMG or UHA-SMG were prepared with different oil volume fractions (60%, 65%, 70%, and 75%, v/v). The macroscopic and microscopic morphology of the resulting HIPEs were then examined and photographed (DM1000, Leica, Germany). Similarly, the appropriate concentration of UHA-SMG was selected from the tested range of 1.0% to 2.0% (w/v).
Confocal laser scanning microscopy (CLSM)
2.6.2
Referring to previous work [24], confocal laser scanning microscopy (LEICA S5, Leica Microsystem, Germany) observation was conducted to reveal the microstructure of the prepared HIPEs stabilized by native SMG or UHA-SMG. Briefly, the HIPEs samples (1 mL) were stained with dyes (20–50 μL) mixed with Nile red (1%, w/v) and fluorescein isothiocyanate isomer (FITC, 1%, w/v). Subsequently, after being placed on a glass slide and covered with a coverslip, 10 μL of stained sample was examined. Images were acquired with excitation at 488 nm and 640 nm.
Morphological observation of HIPEs using light microscopy
2.6.3
A 10 μL aliquot of the fresh HIPEs stabilized by native SMG or UHA-SMG was mounted on a microscope slide and covered with a coverslip. It was then imaged using a Leica DM1000 optical microscope (Germany).
Rheological properties
2.6.4
The rheological behavior of sample was characterized on a DHR-3 rheometer (Waters, USA) equipped with a 40 mm parallel plate (gap: 1000 μm). Frequency sweeps (0.1–10.0 Hz) were performed at a constant strain of 0.2% (within the LVR). Viscosity curves were obtained from flow sweeps at shear rates ranging from 0.01 to 1 s^−1^, with all tests conducted at 25 °C.
Stability analysis of HIPEs stabilized by native SMG and UHA-SMG
2.7
The stability of HIPEs stabilized by native SMG and UHA-SMG was assessed, including thermal stability, centrifugation stability, and storage stability.
Thermal stability
2.7.1
Each HIPE sample (1.5 mL) was aliquoted into a 2.0 mL tube, subjected to heating at 75 °C for 30 min in a water bath, and subsequently cooled to 25 °C. The sample stability was then evaluated for particle size, visual appearance and microscopic morphology.
Centrifugation stability
2.7.2
For centrifugation stability analysis, 1.5 mL of each HIPE was transferred to a 2 mL tube and subjected to centrifugation at 10,000 rpm for 10 min (Sigma high-speed centrifuge 2-16KL, Germany). Prior to imaging, the samples were allowed to settle, and their macroscopic appearance along with droplet characteristics were visually documented.
Storage stability
2.7.3
HIPE samples underwent a 60-day storage test at 4 °C in sealed vials, with subsequent assessment of their visual appearance, micromorphology, and droplet size to determine storage stability.
Encapsulation of lycopene in HIPEs stabilized by UHA-SMG
2.8
For the preparation of LYC-loaded HIPEs, LYC (purchased from Macklin Inc. (Shanghai, China)) was first dissolved in corn oil at a concentration of 1 mg/g and then incorporated into the HIPEs using the method described above. For comparison, a control sample containing LYC dissolved in corn oil was used (CONT). To determine the encapsulation efficiency of LYC, the compound was extracted from the LYC-loaded HIPEs using an n-hexane/ethanol mixture (2:1, v/v). Subsequently, in line with other studies, the content of LYC in the extracts was quantified by measuring absorbance at 472 nm using a UV spectrophotometer (Evolution 220, Thermo Electron, USA) [29], and the retention rate of lycopene was derived with reference to other studies [27].
Stability of lycopene in HIPEs stabilized by UHA-SMG
2.9
UV irradiation stability
2.9.1
The photostability of LYC-loaded HIPEs was assessed by exposing them to a 15 W UV lamp in a closed chamber, followed by quantification of residual LYC at 0, 3, 6, 9, and 12 h. Similarly, the LYC dissolved in corn oil was also treated with a UV lamp (15 W) for varying durations. The LYC retention efficiency was calculated as above.
Thermal stability
2.9.2
Thermal degradation was evaluated by heating the HIPEs in a 75 °C water bath and measuring the residual LYC content at 0, 1, 2, 3, and 4 h.
Storage stability
2.9.3
Storage stability was assessed by storing LYC-loaded HIPEs in light-protected refrigeration at 4 °C [30], followed by LYC quantification after 0, 8, 16, 24, and 32 days.
Statistical analysis
2.10
All the above experiments were replicated at least three times, and the obtained results are expressed as mean ± standard deviation (S.D.). For statistical comparison, one-way ANOVA (analysis of variance) with Tukey's HSD post hoc test (SPSS Statistics 16.0, IBM, USA) was conducted. Statistical significance was set at P < 0.05.
Results and discussion
3
Impact of modification treatment on the particle size, zeta-potential, and solubility of SMG
3.1
Ultrasonic power
3.1.1
As illustrated in Fig. 1A, ultrasonic power significantly influenced the particle size, zeta-potential and solubility of SMG. The native SMG (0 group) exhibited a relatively large particle size (1352.33 ± 31.75 nm), likely due to molecular aggregation resulting from poor solubility. Ultrasonic treatment markedly reduced the particle size of SMG. Specifically, an increase in power from 200 W to 500 W led to a corresponding reduction in average particle size, from 1101.50 ± 27.12 nm to 760.57 ± 34.03 nm. This reduction can be attributed to cavitation-induced effects, high shear forces, and microstreaming generated during ultrasonication, which disrupt non-covalent interactions within and between protein molecules, thereby fragmenting large aggregates [31]. This observed inverse relationship between ultrasonic power and particle size is consistent with findings reported for other plant proteins. For instance, studies on sunflower protein [32] and soy protein [33] isolates have also demonstrated a progressive decrease in particle size with increasing ultrasonic power. Moreover, the zeta-potential of samples was also detected. This analysis provides a direct assessment of a protein's surface charge, a key characteristic influencing its behavior. Elevated charge density serves to amplify repulsive forces between protein molecules, suppressing aggregation and fostering stronger protein-water interactions, thereby boosting overall solubility [34]. The absolute zeta-potential of SMG increased obviously with rising ultrasonic power, indicating a greater net surface charge on the particles. This effect is likely due to the disruption of protein aggregates by high-power ultrasonication, accompanied by a corresponding rise in surface negative charge [35]. The enhanced electrostatic repulsion likely contributes to better dispersibility and stability of SMG. Similar findings have been reported in studies involving Cyperus esculentus seeds [36]. Additionally, a higher absolute zeta-potential generally corresponds to an improved stability of colloidal systems [37]. Protein solubility is a key determinant of its functional properties and, consequently, its potential for industrial applications, as it is governed by the balance of intermolecular forces and modulated by conformational changes [31]. As shown in Fig. 1B, ultrasonic treatment significantly improved the solubility of SMG in a power-dependent manner. The solubility of native SMG was 24.67 ± 0.27%, which increased to a maximum of 31.37 ± 0.47% at 500 W. The turbulence and shear forces generated by ultrasound may disrupt hydrophobic interactions, reducing molecular aggregation and further contributing to the observed increase in solubility [38]. Taken together, the suitable ultrasonic power was selected to 500 W.Fig. 1. Effects of different ultrasonic power (A, B), ultrasonic time (C, D), temperature (E, F), and pH value (G, H) on the particle size, zeta-potential, and solubility of SMG. Values that share a common letter within a row do not differ significantly (P > 0.05).
Ultrasonic time
3.1.2
As can be seen in Fig. 1C and 1D, no significant influence of ultrasonic time in the range of 5–20 min on the particle size, zeta-potential or solubility of SMG was observed. This finding is inconsistent with behaviors reported for myofibrillar protein [39] and tuna protein [40]. As suggested in previous studies, this plateau effect may be attributed to a balance between initial fragmentation and eventual thermally induced aggregation under prolonged ultrasonication. Furthermore, an increase in particle size beyond 30 min of sonication has also been documented in other protein systems, such as β-lactoglobulin [41] and black bean protein isolate [42], supporting the notion that extended treatment can lead to reaggregation. Thus, an ultrasonic time of five minutes was selected for the subsequent experiments.
Modification temperature
3.1.3
As shown in Fig. 1E, as the modification temperature increased progressively from 25 to 100 °C, the particle size of SMG initially decreased, reached a minimum at 70 °C, and subsequently increased. Similar results were found in pea protein isolate [43]. The initial reduction in particle size reflects thermal unfolding and ultrasonic cavitation disrupted the aggregated structures of SMG [44]. The structural changes in the protein induced by heat treatment might result in higher electrostatic repulsion and a weakening of molecular interactions, ultimately enhancing dispersibility [45]. However, compared with 70 °C, the particle size of SMG modified at 90 and 100 °C was both significantly increased (P < 0.05), which may be explained by thermal aggregation of proteins, which is promoted at higher temperatures [46]. As seen from Fig. 1E, the absolute zeta-potential of SMG exhibited a non-monotonic trend with temperature, characterized by an initial rise followed by a decline, which aligns with previous reports [47]. Statistical analysis revealed no significant difference (P > 0.05) in comparisons of the groups treated at 25, 50, and 70 °C. However, once the treatment temperature exceeded 70 °C, a continuous reduction in zeta-potential was observed, with the 100 °C treatment group showing the most pronounced decline. The reduction in zeta-potential during heat treatment can be attributed to protein aggregation. Lee et al. reported that protein matrices formed at 95 °C possessed a more aggregated morphology compared to those formed at 75 °C, leading to charge shielding effects that thereby reduce the net surface charge [48]. Moreover, the conformational changes induced by high temperatures can expose internal cationic residues. The resulting charge neutralization diminishes the net negative surface charge, which is reflected in a lower absolute zeta-potential [47]. Additionally, as illustrated in Fig. 1F, SMG solubility showed an initial rise followed by a decline, a trend that may be closely associated with the dynamic changes in protein particle size [49].
pH value
3.1.4
As shown in Fig. 1G and 1H, when the pH was 5.0, the particle size, zeta-potential, and solubility of SMG all exhibited inferior performance. The maximum particle size observed at pH 5.0 is likely due to the proximity of the protein to its isoelectric point. However, deviation of pH from the isoelectric point results in increased electrostatic repulsion, thereby inducing the a corresponding reduction in particle size [50]. Moreover, as the pH rose from 3.0 to 5.0, the zeta-potential transitioned from positive to negative. The low absolute zeta-potential under acidic conditions suggests weakened electrostatic repulsion, which allowed particle aggregation, ultimately leading to reduced solubility [51]. Similar phenomena have also been reported in studies on Chenopodium seed protein isolate [52]. As the pH further increased, zeta-potential and solubility of SMG both showed gradual improvement. This phenomenon can be explained by the unfolding and refolding of protein molecules at higher pH, which disrupts hydrophobic and van der Waals interactions, thereby enhancing solubility [49]. Moreover, this observed increase in protein solubility likely results from stronger inter-protein electrostatic repulsion coupled with an enhanced capacity for hydrogen bonding with water [53]. However, considering the potential adverse effects of a highly alkaline environment on subsequent experiments, pH 9.0 was ultimately selected as the optimal pH.
Difference in the structure properties between native SMG and UHA-SMG
3.2
Far-UV circular dichroism (CD) spectra
3.2.1
An established technique in protein science, far-UV circular dichroism detection was employed to characterize the secondary structure composition, folding, and binding properties of samples. As presented in Fig. 2A, distinct spectral differences were observed between the native SMG and UHA-SMG. The presence of a positive peak near 190 nm and a negative peak near 208 nm suggests that α-helix is the predominant secondary structure in UHA-SMG [54]. Following modification, an increase in the negative peak intensity at 208 nm was observed in UHA-SMG. Moreover, it can be observed from Fig. 2B that the secondary structure composition of SMG exhibited significant changes following modification, most notably an increase in α–helix content. Upon modification, an increase in α-helix content (from 20.33 ± 0.32% to 50.77 ± 0.93%) was accompanied by a decrease in the proportions of β-sheet, β-turn, and random coil. The sharp increase in α-helix content may be partially attributed to the effects of high-intensity ultrasound, which is in agreement with the work of Hu et al., whereas high ultrasonic power has been shown to increase α-helix content [55]. Additionally, it has been reported that both alkaline pH and elevated temperature treatments can increase the α-helix content of certain proteins. For example, the α-helix ratio of egg white protein was also found to increase after alkaline pH treatment (pH = 11) [56]. As reported by Wang et al., heat treatment led to a rise in α-helix content of soybean protein, accompanied by a decrease in β-sheet content compared to the untreated sample [57]. In addition, these findings are not consistent with reported changes in the secondary structure of lentil proteins treated by pH-shifting and heating, where a decrease in α-helix and an increase in β-sheet content were observed relative to the native protein [58]. In protein secondary structure, intra-chain hydrogen bonding characterizes the α-helix, whereas inter-chain hydrogen bonding facilitates β-sheet assembly [59]. The elevated α-helix content drove the development of multilayer surface interfaces, which in turn enhanced noncovalent interactions-both within the protein and with surrounding water molecules-leading to superior hydration properties [60]. The partial loss of β-sheet structure, which is typically buried within the protein interior, suggests the exposure of hydrophobic sites. This exposure can thereby lead to an increase in surface hydrophobicity [57]. This inference was further confirmed by subsequent experimental measurements of surface hydrophobicity. These findings indicate that the modification might disrupt the network stabilizing the original β-sheets, β-turns, and random coils of SMG, thereby allowing the polypeptide chain to refold into an overall more stable protein conformation. Contrasting with the results of this work, other research has demonstrated different outcomes. A study by Xie et al., for example, found an increase in both α-helix and β-sheet contents of myofibrillar protein after ultrasound-assisted first-stage thermal treatment [59]. The alterations in protein secondary structure induced by modification are likely due to multiple factors, including ultrasonic duration and the intrinsic properties of the protein, which collectively account for the observed discrepancies [61].Fig. 2. Comparative analysis of the structural properties between SMG and UHA-SMG. (A) Far-UV circular dichroism spectra. (B) Secondary structure composition. (C) UV–Vis absorption spectra. (D) Fluorescence spectra. (E) Surface hydrophobicity analysis of SMG. (F) Surface hydrophobicity analysis of UHA-SMG.
UV-Vis spectroscopy
3.2.2
UV–Vis spectroscopy was employed to monitor changes in the tertiary structure of samples by detecting the absorption spectra of their intrinsic aromatic amino acids, whose spectral properties are sensitive to local microenvironment [62]. As illustrated in Fig. 2C, both native SMG and UHA-SMG exhibited a characteristic absorption peak at approximately 280 nm, which may be related to the π → π* electronic transitions of aromatic amino acids (phenylalanine, tryptophan, and tyrosine) [63]. Notably, the intensity of this maximum absorption peak of UHA-SMG was slightly lower than that in native protein. This decrease in absorbance intensity may result from structural alterations induced by modification. This phenomenon has been similarly reported in studies involving other plant proteins; for instance, both rice protein and zein have shown analogous spectral changes upon modification [28].
Fluorescence spectroscopy
3.2.3
Fluorescence spectroscopy of native SMG and UHA-SMG was measured in this study to further monitor changes in the tertiary structure of proteins [64]. Alterations in the intrinsic fluorescence spectra, such as shifts in the emission wavelength or changes in intensity, reflect modifications in the microenvironment of aromatic residues (e.g., tryptophan and phenylalanine), which are indicative of protein conformational changes [65]. As depicted in Fig. 2D, the UHA-SMG exhibits higher fluorescence intensity than the native SMG, suggesting that tryptophan residues and hydrophobic groups are more likely to be located in a more hydrophilic microenvironment under modified conditions. Previous studies indicate that ultrasound-induced protein depolymerization exposes previously buried chromogenic groups, rendering them accessible to the aqueous solvent [60]. Moreover, the unfolding of protein structures induced by heat or pH-shifting also contributes to this phenomenon, as it exposes previously buried hydrophobic groups. This observation is consistent with findings reported for rice glutelin [66], donkey milk whey protein [67], and egg yolk protein [68].
Surface hydrophobicity
3.2.4
Since the exposure of hydrophobic groups reflects protein conformational changes and directly affects functional properties, determining its extent is critical [69]. Therefore, to assess the extent of hydrophobic group exposure on protein surfaces of native SMG and UHA-SMG, the H0 of two samples was determined. As illustrated in Fig. 2E and F, the combined modification treatment significantly enhanced the H0 of SMG, indicating that the treatment effectively promoted protein unfolding, leading to greater exposure of internal hydrophobic regions. The primary mechanism involves the cavitation-induced mechanical shear and perturbation from ultrasonication, which facilitates structural expansion and the exposure of hydrophobic residues [24]. Additionally, the higher surface hydrophobicity was also found following alkaline treatment in previous work. This was due to the pH-induced stretching of protein molecules and the subsequent inability to fully refold upon neutralization, which left hydrophobic groups exposed [68]. Concurrently, the accompanying thermal and alkaline treatments further intensified protein denaturation, disrupting the native conformation and collectively promoting the exposure of additional hydrophobic residues, thereby elevating the overall H0. However, in general, higher surface hydrophobicity in proteins tends to be accompanied by lower solubility, which was inconsistent with our results. It is hypothesized that factors other than hydrophobicity may influence the solubility of SMG. Previous studies have indicated that protein solubility can also be governed by spatial conformation and intermolecular interactions [68]. This finding aligns with the results of previous studies [53], [70]. These results demonstrate that ultrasonic-heat-pH treatment effectively modifies the tertiary conformational state of SMG molecules.
Comparative analysis of the microscopic morphology between native SMG and UHA-SMG
3.3
The micromorphological features of native SMG and modified UHA-SMG were shown in Fig. 3A. Clearly, significant alterations in the morphology of SMG were observed after modification treatment. It can be seen that the native SMG exhibited irregular, flocculent aggregates, whereas the UHA-SMG displayed a smoother surface with a porous structure, appearing more compact and dense overall. This morphological transformation is likely due to the structural reorganization of the protein surface induced by treatment, which aligns with findings reported for rice protein [28]. The synergistic action of shear forces, impact effects, and cavitation generated by ultrasonication combined with thermal treatment and pH-shifting collectively contributed to the dispersion and reorganization of protein aggregates.Fig. 3. Comparative evaluation of ultrasound-heating-pH-shifting modification on the properties of SMG. (A) Microscopic morphology. (B) Contact angle. (C) Interfacial tension.
Interface properties of native SMG and UHA-SMG
3.4
The three-phase contact angle (θ) serves as a fundamental parameter for assessing the wettability of solid particles [71]. Generally, a θ of 90° is regarded as the optimum condition for the adsorption of particles at a fluid–fluid interface. Fig. 3B displays the contact angles of SMG before and after modification. The native SMG exhibited a contact angle of 44.87 ± 1.68°, which reflects low interfacial affinity and weak emulsion stabilization. The increased contact angle (85.40 ± 0.62°) of SMG post-modification indicated enhanced interfacial activity and adsorption, which are conducive to improved emulsion stability. Similar results were also found in other proteins (rice protein [28], Tenebrio molitor proteins [72], copra meal protein [73], bamboo fungus protein [74]) modified by ultrasonic, higher pH, or heating. As an example, an increase in the θ of pea protein isolate-tannic acid complexes following ultrasonic treatment was observed by Yu et al. [75]. The observed differences in contact angle may arise from structural alterations in the protein molecules induced by the combined treatment of ultrasound, heating, and pH-shifting [75]. Furthermore, the contact angle also reflects the hydrophilic/hydrophobic character of the particles [74]. Clearly, the UHA-SMG exhibited stronger hydrophobicity, which is consistent with the results mentioned above. This enhanced intermediate wettability indicates that the UHA-SMG is more suitable as a stabilizer for emulsions.
Protein adsorption at the oil–water interface serves as a key indicator of emulsion stabilization potential [76]. A complete and saturated interfacial layer is essential for emulsion stability, as it directly acts as a barrier against the coalescence and flocculation of oil droplets [77]. A greater interfacial tension exhibited by proteins at the oil–water interface increases the energy needed to disrupt the interface during emulsification, which in turn impairs emulsion stability [78]. Therefore, the dynamic interfacial tension measurements were performed on the two samples. This technique is commonly used for examining protein adsorption at the oil–water interface [79]. As depicted in Fig. 3C, both samples exhibited a time-dependent decline in oil/water interfacial tension, resulting from the continuous adsorption of proteins or protein complexes at the interface [80]. Furthermore, the time-dependent interfacial tension profile revealed a two-stage adsorption pattern for SMG. The initial steep decline (0–1800 s) corresponded to the rapid attachment of SMG molecules to the oil–water interface. Upon further extension of time, the decrease in interfacial tension gradually slowed and eventually almost stabilized, indicating that interfacial adsorption gradually reached saturation. Furthermore, the interfacial tension of UHA-SMG was obviously lower than that of native SMG. The drops of the modified protein solution appeared more elongated at equilibrium, consistent with a lower surface tension. The reduced interfacial tension of UHA-SMG might be ascribed to the enhanced solubility of SMG following ultrasonic-alkaline-thermal treatment, which bolsters the protein's binding capacity at the interface and thus brings about a marked reduction in interfacial tension. Additionally, the reduction in interfacial tension observed for UHA-SMG can also be attributed to their increased surface hydrophobicity, which drives more efficient adsorption at the interface. Meanwhile, the small particle size and higher absolute zeta-potential of UHA-SMG facilitate rapid diffusion to the oil–water interface, while the exposure of hydrophobic groups lowers the kinetic barrier for adsorption, promoting efficient interfacial occupancy [79], [81]. A similar decrease in interfacial tension was also observed upon ultrasonication, pH-shifting, thermal treatment in other proteins (mussel actomyosin [82], protein from the shrimp heads [83], soy protein [84], etc.).
Formation and morphology of HIPEs stabilized by UHA-SMG
3.5
The microscopic and visual appearance of emulsions stabilized by native SMG (2%) or UHA-SMG (1.0%, 1.5%, 2%) were observed at different corn oil amounts (internal phase volume, 60%, 65%, 70%, and 75%). The microscopic morphology of emulsions stabilized by native and UHA-SMG exhibited distinct differences, as illustrated in optical microscopy results (Fig. 4). No uniform emulsion droplets were observed in any formulations prepared with 2% native SMG, regardless of the oil phase amount. Moreover, these formulations also failed to form gel-like emulsions (Fig. 4B). In contrast, gel-like emulsions were obtained with 1% UHA-SMG in all groups except at an oil phase volume of 60%. Furthermore, the influence of UHA-SMG concentration on HIPEs formation was evaluated. Gel-like emulsions stabilized by 1.5% UHA-SMG were established at corn oil amounts of 65%, 70% and 75%, whereas those prepared with 2.0% UHA-SMG exhibited a uniform and self-supporting appearance in all oil phase volumes used. Optical microscopy further revealed that emulsions stabilized by 2.0% UHA-SMG possessed more regularly shaped droplets and a densely packed microstructure. Furthermore, the images show that the HIPEs formed under these conditions completely sedimented within the glass vial. Importantly, the sedimented phase exhibited no flow upon inversion of the vial, which is a signature property of high internal phase emulsions. Previous research indicates that in protein-stabilized emulsion systems, increasing the concentration of protein particles enhances the surface charge density of the droplets. This elevated charge promotes stronger electrostatic repulsion among droplets, thereby suppressing their aggregation and consequently improving emulsion stability [85]. Moreover, studies have reported that increasing the protein particle concentration can markedly improve emulsion stability by promoting the formation of a more extensive stabilizing network and increasing protein coverage on the oil droplet surface [52], [85]. Overall, these results indicate that, compared with its native counterpart, the UHA-SMG is more suitable for stabilizing emulsion. Additionally, as mentioned above, the minimum threshold of the internal phase volume fraction in HIPEs was 74%. Thus, the HIPEs stabilized by 2% UHA-SMG at an internal oil volume of 75% was selected for subsequent further analysis.Fig. 4. Effect of UHA-SMG concentration and corn oil content on the microstructure (A) and macroscopic appearance (B) of emulsions.
Microstructure and rheological characterization of HIPEs stabilized by UHA-SMG
3.6
The microstructure of emulsions stabilized by UHA-SMG and SMG was detected using confocal laser scanning microscopy. In this micrograph, the red signal indicates the inner oil phase of the droplets, whereas the green signal corresponds to the surrounding water phase (Fig. 5A). Clearly, the system stabilized with 2% (w/v) SMG showed no visible droplet microstructure but exhibited evident oil droplet aggregation under microscopy. This suggests that the protein interfacial layer was not strong enough to prevent droplet flocculation and physical compression during sample preparation. Conversely, at 1% (w/v) protein, UHA-SMG formed more regular spherical droplets, although its interfacial film ruptured under external pressure. This indicates that the combined modification improved the interfacial activity of SMG, yet the resulting film still lacked sufficient interfacial coverage or mechanical strength at this concentration. However, as the UHA-SMG concentration increased to 1.5% or 2% (w/v), the emulsion systems consisted of red spheres embedded within a green matrix, suggesting the formation of O/W HIPEs. Moreover, the spherical morphology of emulsion droplets is likely due to the generation of a compact interfacial film by UHA-SMG. This film envelops the droplets within the continuous phase, thereby effectively suppressing deformation, collision, flocculation, and coalescence [86]. These results demonstrate that a sufficient concentration of UHA-SMG ensured complete coverage of the oil droplet interfaces and facilitated the formation of a high-strength viscoelastic interfacial film.Fig. 5. Microstructure and rheological properties of HIPEs stabilized by SMG and UHA-SMG. (A) Confocal laser scanning microscopy detection. (B) Frequency sweep profiles (storage modulus, G'; loss modulus, G''). (C) Apparent viscosity as a function of shear rate.
The physical stability of the prepared formulation is largely governed by its rheological behavior, primarily reflected in apparent viscosity and viscoelasticity. Emulsion stability is substantially improved by increased apparent viscosity, which effectively inhibits droplet Brownian motion, collision, and aggregation [87]. Fig. 5B depicts the results of the dynamic frequency sweep measurements, where the storage modulus (G') and loss modulus (G'') characterize the solid-like elastic and liquid-like viscous responses of the system, respectively [88]. Regardless of protein concentration, both G' and G'' of UHA-SMG-stabilized HIPEs increased with frequency, implying that the observed rigidity originated from a physically cross-linked network between droplets [89]. Moreover, in all samples, the G' was significantly greater than the G''. This demonstrates that the prepared HIPEs behaved as elastic solids rather than viscous fluids throughout the entire frequency sweep [90]. Notably, the HIPEs stabilized by 2% (w/v) UHA-SMG exhibited obviously higher G' and G'' values than other groups. This superior viscoelasticity results from a more compact and cohesive microstructure, which in turn confers greater resistance to deformation. Furthermore, it is noteworthy that both the G' and G'' increased with elevating the UHA-SMG concentration, indicating that elevating the UHA-SMG content enhances the viscoelasticity of the HIPEs. This result aligns well with the findings of previous research, which suggests that the performance might be attributed to the reduced droplet size at higher protein concentrations, contributing to a more compact and densely packed organization of the HIPEs. This structural compactness subsequently strengthened the network's ability to resist deformation and droplet coalescence.
As shown in Fig. 5C, all UHA-SMG-stabilized HIPEs displayed characteristic shear-thinning behavior, where apparent viscosity decreased with increasing shear rate, confirming their pseudoplastic nature [91]. The initial transient rise in apparent viscosity at the beginning of the measurement may be attributed to insufficient equilibration at the interface between the measuring fixture and the sample [24]. As shear intensity increased, the aggregated structure of oil droplets within the emulsion gradually broke down, leading to a reduction in flow resistance and macroscopically manifested as a decrease in viscosity [92]. Furthermore, as the concentration of UHA-SMG increased, the emulsion viscosity rose markedly, which is primarily due to the higher protein concentration leading to the formation of smaller and more numerous oil droplets, thereby enhancing spatial interactions and structural compactness among droplets [90]. The denser droplet arrangement and stronger interfacial film structure collectively endowed the system with higher apparent viscosity and greater resistance to deformation, consequently improving its physical stability.
Stability analysis of HIPEs stabilized by UHA-SMG
3.7
Centrifugal stability
3.7.1
Given that the centrifugal stability of emulsions can serve as an indicator of their resistance to gravity during storage, this study evaluated the centrifugal stability of UHA-SMG-stabilized HIPEs (Fig. 6). It was observed that all HIPE samples underwent water phase separation upon centrifugation at 10,000 rpm (Fig. 6A). Additionally, obvious oil loss (top layer) was found in emulsions stabilized by SMG, indicating its poor centrifugal stability. In contrast, the incorporation of UHA-SMG led to a marked improvement in the stability of the HIPEs. It is worthy noting that increasing the UHA-SMG concentration from 1.0% (w/v) to 2.0% (w/v) yielded a gradual decrease in the precipitated aqueous phase as well as oil loss. For the formulations containing 1.5% and 2.0% UHA-SMG, only minimal water and negligible oil were separated after centrifugation, suggesting the adsorbed protein layer on oil droplets effectively suppressed flocculation. Additionally, the enhanced centrifugal stability of HIPEs with high UHA-SMG content is also likely related to their high viscosity, viscoelasticity, and strong gel network, which minimized droplet–droplet collisions under high centrifugal force, thereby preventing aggregation [93], [94]. Moreover, the lower oil loss observed in HIPEs stabilized by 2.0% UHA-SMG can be attributed to their smaller average droplet size, which is also supported by the particle size measurements result (Fig. 6B and C) [95]. As depicted in Fig. 6B, after centrifugation, no obvious change in particle size was observed in the samples incorporated with 1.5% (w/v) and 2.0% (w/v) UHA-SMG (P > 0.05).Fig. 6. Evaluation of the physical stability of UHA-SMG-based HIPEs under stress treatments. (A–C) Centrifugation. (D–F) Heat treatment. (G–I) Storage for two months. The panels in each column show the visual appearance, droplet size distribution, and microscopic morphology, respectively. Statistical significance is denoted as *P < 0.05, **P < 0.01, and ***P < 0.001.
Thermal stability
3.7.2
Since thermal treatment is common in food processing and consumption, the HIPEs prepared with SMG or UHA-SMG were subjected to a 75 °C/30 min heat treatment. Subsequently, their droplet size and appearance were recorded. As depicted in Fig. 6D and E, after heating, obvious oil leakage along with a notable enlargement in droplet size were observed in the emulsions stabilized with SMG (2%) or UHA-SMG (1% and 1.5%) (P < 0.05). Specifically, the droplet size in the emulsion stabilized with 2% SMG increased from an initial 64.90 ± 1.55 μm to 103.93 ± 1.63 μm, while that in the emulsion stabilized with 1% UHA-SMG increased from 29.88 ± 0.64 μm to 39.10 ± 1.19 μm (Fig. 6E). The observed increase in droplet size suggests thermal-induced coalescence of emulsion droplets. Optical microscopy further verified droplet aggregation within these HIPEs (Fig. 6F). Remarkably, nearly no oil droplets were identified in the 2% SMG-stabilized samples. In contrast, the HIPEs prepared with 1.5% UHA-SMG exhibited only minimal water separation at the bottom and an increase in droplet size (from initial 22.75 ± 1.25 μm to 24.97 ± 1.61 μm). On the contrary, as illustrated in Fig. 6D, the appearance of HIPEs stabilized by 2.0% UHA-SMG did not change significantly after heating. This behavior can be explained by the formation of a more stable interfacial layer at higher UHA-SMG concentrations, which facilitate the development of a resilient, gel-like network structure [96].
Storage stability
3.7.3
As a thermodynamically unstable system, HIPEs are susceptible to destabilization during long-term storage, including demulsification, oil-phase sedimentation, and fat floating [27]. In this study, the storage stability of different HIPE systems was systematically evaluated (Fig. 6G–I). Unlike the emulsions stabilized with 2% (w/v) SMG or 1% (w/v) UHA-SMG, those prepared with 1.5% or 2.0% UHA-SMG showed no macroscopic oil separation or creaming during the 60-day storage (Fig. 6G). These two samples maintained homogeneity and stability in vials, with no flow upon inversion. Furthermore, the particle size analysis results showed that after storage, no statistically significant difference was found in the 1.5% (w/v) and 2.0% (w/v) UHA-SMG groups (P > 0.05). These findings reveal the storage stability of HIPEs stabilized by 2.0% (w/v) UHA-SMG. This property might be related to the smaller droplets stabilized by UHA-SMG.
Taken together, the above findings demonstrate that the HIPEs incorporated with 2.0% (w/v) UHA-SMG exhibited excellent centrifugal, thermal, and storage stability. Meanwhile, these results also demonstrate that the combined modifications involving ultrasonication, heating, and pH-shifting effectively led to significant enhancements in both the gel strength and storage stability of SMG-based HIPEs.
Encapsulation and stability of lycopene in HIPEs stabilized UHA-SMG
3.8
Encapsulation of lycopene in HIPEs
3.8.1
Lycopene (LYC), a liposoluble carotenoid from tomatoes, offers significant health benefits, including cardiovascular and cancer prevention. However, its poor stability, solubility, and bioavailability limit its application [97]. Therefore, the encapsulation of LYC is of critical importance. Based on the good stability of the HIPEs prepared here, this study explores its application for encapsulating LYC. As shown in Fig. 7A, the emulsion droplets loaded with LYC exhibited an orange-red color and maintained a regular spherical shape in UHA-SMG-stabilized HIPEs. All the HIPEs exhibited a comparable orange color, and the self-supporting property of those stabilized by 1.5% or 2% UHA-SMG remained unchanged after LYC encapsulation (Fig. 7B). These observations suggest that the UHA-SMG-based HIPE system holds promise as an effective delivery vehicle for LYC. To evaluate the quality and application potential of the HIPE systems, the encapsulation efficiency of LYC was determined. Notably, as presented in Fig. 7C, the LYC encapsulation efficiency demonstrated a marked increase at higher protein concentrations, reaching 95.38 ± 1.65% at 1.5% (w/v) and 96.39 ± 1.34% at 2.0% (w/v), which were significantly greater than the efficiencies observed in the remaining two groups (P < 0.05), also confirming the effective encapsulation ability of HIPEs stabilized by 1.5% and 2.0% UHA-SMG. Additionally, compared to the pseudo-solid appearance of the 1% UHA-SMG HIPEs described earlier, the LYC-loaded HIPEs stabilized by 1% UHA-SMG exhibited a marked increase in fluidity. This phenomenon is likely due to LYC hydrophobicity raising the oil phase viscosity and lowering emulsification efficiency, leading to droplet coarsening and ultimately resulting in an intensified phase separation tendency [98].Fig. 7. Encapsulation efficiency and stability of lycopene in UHA-SMG-stabilized HIPEs. (A) Representative microscopy image of lycopene-loaded HIPEs. (B) Macroscopic appearance of lycopene-encapsulated HIPEs. (C) Lycopene encapsulation efficiency of prepared HIPEs. (D–F) Stability of encapsulated lycopene under UV irradiation (D), thermal treatment (E), storage (F). Data marked with the same letter within each row are not significantly different (P > 0.05).
Stability assessment of lycopene in HIPEs
3.8.2
LYC is highly susceptible to degradation during manufacturing and storage, especially under intense light and high temperature [99]. Extensive evidence reveals that exposure to UV light or high temperature can induce the isomerization of bioactive all-trans-lycopene to various cis isomers, which exhibit reduced or negligible bioactivity [100]. Therefore, the UV-protective effect and thermal stability of UHA-SMG-stabilized HIPEs were investigated, respectively. While all samples showed a gradual decline in retention rate of LYC with irradiation time, the 2.0% UHA-SMG-stabilized HIPEs maintained higher levels (Fig. 7D). Notably, free lycopene in corn oil degraded rapidly. After 12 h of irradiation, the retention rate of lycopene in the CONT group was only 58.56 ± 0.23%, whereas the values were higher in HIPEs stabilized by native 2.0% SMG, 1.0% UHA-SMG, 1.5% UHA-SMG, and 2.0% UHA-SMG, with values of 65.13 ± 1.94%, 72.05 ± 2.07%, 80.20 ± 1.39%, and 81.71 ± 0.29%, respectively. This enhanced stability is attributed to the encapsulation of lycopene within the HIPE's internal oil phase, which not only prevents its direct exposure to UV light through physical shielding but also likely hinders the penetration of pro-oxidative species to the core [30], [100].
To ensure microbial safety and extend shelf life, heat treatment is a common practice across food processing, packaging, and storage operations. Therefore, improving the stability of LYC during heating is of critical importance. As depicted in Fig. 7E, the retention rate of LYC in all groups gradually decreased as heating time increased from 0 to 4 h. However, after four hours of treatment, the LYC retention in all HIPE systems was significantly higher than in the CONT group (P < 0.05), indicating that HIPEs effectively mitigate lycopene's thermal degradation. This improvement is attributed to the encapsulation of LYC within the oil droplets, which are coated by a dense UHA-SMG interfacial layer. This physical barrier not only restricts the penetration of heat and oxygen into the droplets but also prevents droplet coalescence and shields LYC from oxidative species generated at high temperatures [98].
The stability of the HIPEs loaded with LYC was further evaluated over a 32-day storage period at 4 °C, which is shown in Fig. 7F. All HIPE formulations provided significantly better protection for lycopene than corn oil. Further comparison showed that while the emulsion stabilized with 2.0% SMG conferred initial protection, its retention rate declined sharply and ultimately approached that of corn oil. This sharp decline indicates that the interfacial layer formed by native SMG lacked the long-term stability needed to effectively block oxygen and light-induced degradation. In contrast, the emulsion with 1.0% UHA-SMG maintained significantly higher lycopene retention than the SMG throughout storage. This result directly demonstrates that the ultrasound–heating–pH-shifting modification effectively enhanced the interfacial stability and barrier properties of the protein. Additionally, the protective effect improved in a stepwise manner as the concentration of UHA-SMG increased from 1.0% to 2.0%. Among these, the HIPEs stabilized with 2.0% UHA-SMG exhibited the highest lycopene retention rate (81.16% ± 2.28%) after 32 days of storage, showing marked differences relative to all other groups (P < 0.05).
Conclusions
4
In the present study, the synergistic ultrasound-heating-pH-shifting treatment was demonstrated to effectively enhance the physicochemical properties and emulsifying functionality of SMG. This process disrupted the native protein aggregates, leading to reduced particle size, lower interfacial tension, altered micromorphology, and enhanced functional properties, including solubility, zeta-potential, interfacial activity, and surface hydrophobicity. Structural analysis further indicated that the modification altered both the secondary and tertiary structure of SMG. At 2.0% concentration, the UHA-SMG successfully stabilized oil-in-water HIPEs, which exhibited a dense droplet structure, shear-thinning behavior, solid-like viscoelasticity, and excellent stability. Furthermore, the HIPEs achieved high lycopene encapsulation efficiency (96.39 ± 1.34%) and significantly retarded its degradation under UV light, heat, and storage, owing to the protective gel network and dense interfacial layer. Collectively, this study confirms that ultrasound-heating-pH-shifting modification is an efficient strategy to develop SMG as a bio-based stabilizer for lipophilic bioactive delivery systems, while also offering a promising strategy for the comprehensive valorization of safflower seed meal, contributing to the sustainable utilization of agricultural by-products. However, further research is needed to evaluate the in vivo bioactivity of the encapsulated lycopene and to explore the feasibility of large-scale modification of the target proteins. Moreover, the application of UHA-SMG-stabilized HIPEs for delivering other bioactives is also worth exploring in future work.
Consent for publication
5
Not applicable.
CRediT authorship contribution statement
Ke'er Xiao: Writing – original draft, Methodology, Investigation, Formal analysis. Lingyu Gao: Visualization, Methodology, Investigation, Data curation. Qiuyu Lu: Validation, Methodology. Qiaoyu Wang: Visualization, Methodology, Investigation, Data curation. Ziteng Zhao: Validation, Methodology. Mukaddas Sai: Validation, Methodology. Xinyu Meng: Validation, Methodology. Lili Guan: Writing – review & editing. Jing Yang: Writing – review & editing. Linna Du: Writing – review & editing, Project administration, Funding acquisition, Data curation, Conceptualization.
Ethics approval and consent to participate
Not applicable.
Funding
This research was funded by Jilin Province Science and Technology Support Program of P.R. China (grant number, 20250102294JC); Xinjiang safflower industry development fund.
Declaration of competing interest
The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.
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