Study on stabilized mechanism of high internal phase Pickering emulsions based on commercial yeast proteins: Modulating the characteristics of Pickering particle via sonication

Graphical abstract

Highlights

• •The sonication modulated the structure and interface characteristics of YPs.
• •The HIPPEs was successfully characterized using the YPs particles for the first time.
• •The stabilization mechanism of YPs on HIPPEs varies depending on the sonication time.
• •The present results shed new light on the application of commercial YPs.

Abstract

The primary significance of this work is that the commercial yeast proteins particles were successfully used to characterize the high internal phase Pickering emulsions (HIPPEs). The different sonication time (0,3,7,11,15 min) was used to modulate the structure and interface characteristics of yeast proteins (YPs) that as Pickering particles. Immediately afterward, the influence of YPs particles prepared at different sonication time on the rheological behavior and coalescence mechanism of HIPPEs was investigated. The results indicate that the YPs sonicated for 7 min exhibited a more relaxed molecular structures and conformation, the smallest particle size, the highest H₀ and optimal amphiphilicity (the three-phase contact (θ) was 88.91°). The transition from extended to compact conformations of YPs occurred when the sonication time exceeded 7 min, resulting in an augmentation of size of YPs particles, a reduction in surface hydrophobicity (H₀), and an elevation in hydrophilicity. The HIPPEs stabilized by YPs particles sonicated for 7 min exhibited the highest adsorption interface protein percentage and a more homogeneous three-dimensional (3D) protein network, resulting in the smallest droplet size and the highest storage (G′). The HIPPEs sample that stabilized by YPs particles sonicated for 15 min showed the lowest adsorption protein percentage. This caused a reduction in the thickness of its interface protein layer and an enlargement in the droplet diameter (D _[3,2]). It was prone to droplet coalescence according to the equation used to evaluate the coalescence probability of droplets (Eq (2)). And the non-adsorbed YPs particles form larger aggregation structures in the continuous phase and act as “structural agents” in 3D protein network. Therefore, mechanistically, the interface protein layer formed by YPs particles sonicated 7 min contributed more to HIPPEs stability. Whereas the “structural agents” contributed more to HIPPEs stability when the sonication time exceeded 7 min. The present results shed important new light on the application of commercial YPs in the functional food fields, acting as an available and effective alternative protein.

1. Introduction

High internal phase emulsions (HIPEs), a type of emulsion with disperse phase volume fractions is greater than 74.05 % [1]. They were widely used in some nutritionally fortified functional food products, such as sauces, spreads and salad dressings [2]. Due to their tunable semisolid behavior, smooth textures in the mouth, and ability to hold large amounts of fat-soluble bioactive substances [3]. In comparison to HIPEs stabilized by traditional surfactants, high internal phase Pickering emulsions (HIPPEs) offer numerous benefits such as reduced stabilizer requirement, enhanced resistance to coalescence, improved storage stability, and decreased environmental pollution [4]. The involvement of Pickering particles in the emulsion framework formation has captured the attention of researchers, making it a significant area of research. In the past few years, for instance, researchers have investigated the utilization of inorganic and organic particles in the fields of food and medicine [5], [7]. Nevertheless, the utilization of inorganic particles is restricted because of concerns regarding the environment and the safety of food. Newer research for Pickering emulsions has primarily concentrated on Pickering particles made from edible proteins in the field of food, because of their amphiphilic properties, minimal toxicity/non-toxic, and eco-friendliness [8]. Currently, these proteins primarily derived from animal and plant, including those constructed from whey protein isolate [9], myofibrillar protein [10], quinoa protein [11], zein [12], gliadin [15], pea protein [16], kafirin [17], and soy protein [6]. There have been relatively few studies on the protein-based Pickering particles of microbial sources.

The increasing yearly per capita protein demand and global protein consumption since the beginning of the century have led to a surge in research interest in the development of sustainable alternative proteins [18], [19]. As a microbial protein, the yeast proteins (YPs) are one of the most desirable new alternative proteins. This stems from its low allergenicity (almost allergen free), sustainability, odorless, and economical [18]. Typically, they are acquired from yeast cells that are cultured in inexpensive waste from the food industry, as well as waste from forestry and agriculture. Spent brewer's yeast, for instance, the protein account for 40 %–60 % of its dry weight [21]. Which was one of the predominant by-products in beer brewing process [22]. In parallel, proteins derived from yeast are considered to be a superior source of complete protein due to their comprehensive range of amino acids and amino acid composition that closely resembles the optimal protein level [23]. However, the isolation and extraction techniques of industry caused a compact “closed” conformation of commercial YPs, thereby substantially diminished its physiological functions and downstream processing characteristics (such as solubility, emulsifying property, and digestibility) [24], [25]. This also limits its potential uses as a new type of edible protein resource. As a matter of fact, there were relatively few research articles on the application of YPs as alternative proteins. Yang et al. [26] utilized YPs-chitooligosaccharide composite to stabilize the bioactive molecule to stabilize the bioactive molecule–betanin. Wang et al., [28] developed a novel edible scaffold for the production of high-quality cell-cultured meat by partially replacing collagen with YPs. Guo et al. [29] prepared emulsified sausages using YPs as an animal fat replacer. Xia et al. [20] prepared meat analogues were prepared using YPs and konjac glucomannan blends. Thus, improving the processing adaptability of commercial YPs was essential for expanding its application fields and utilization. Typically, many commercial proteins, especially those without suitable particle wettability/surface charge, or low interfacial activity, exhibited poor coalescence stability [30]. The application of ultrasonic cavitation can destroy the secondary, tertiary and quaternary structure of proteins by disrupt the non-covalent bonds, thereby altering the wettability of protein particles, without any significant change on their primary structure [31], [32]. This is because in food industries, the sonication has high overall energy efficiency (80–90 %) to convert electricity into mechanical forces to generate numerous stresses onto the protein particles [33].

The purpose of the work was to improve the interface performance of the commercial YPs in the HIPPEs field and made it an available and effective alternative protein. Using different sonication time to modulate the structure and interfacial properties of YPs as Pickering particles. Including its conformation, secondary structure, particle size, ζ-potential, surface hydrophobicity, amphiphilicity, and interfacial tension. Furthermore, the HIPPEs was successfully characterized using the YPs particles for the first time, and the influence of YPs particles prepared at different sonication time on the coalescence mechanism of the HIPPEs was investigated. Which is mainly based on the droplet size, microstructure, percentage of adsorbed protein, rheological behavior and the equation used to evaluate the coalescence probability of droplets. This work offered a new direction for future research of commercial YPs, which may be of practical application for food industry.

2. Materials and methods

2.1. Materials and chemicals

Yeast proteins (YPs) (type: F81, protein content is 81.1 % on a dry basis) was supplied by Angel Yeast Co., Ltd. (Wuhan, China). Grade 1 soybean oil was supplied by Jiusan Grain and Oil Industry Group Limited (Harbin, China). All chemicals were of laboratory analytical reagent grade, which were purchased from Sinopharm Chemical Reagent Co., Ltd. (Shanghai, China) and used without further purification.

2.2. Preparation of YPs particles

The YPs particles were fabricated using the method of Qin, Luo, and Peng [34] with a few modifications. The YPs powder was dispersed into phosphate buffer (pH 7.0) to form a concentration of 5 % (w/t) dispersion liquid. The dispersion was stirred in a magnetic agitator for 2 h, and then put in the refrigerator (4 °C) overnight. Finally, 30 ml YPs suspensions were sonicated under 3 min, 7 min, 11 min and 15 min (JY98-IIIN, Xinzhi Technology Co., Ltd., Ningbo, China), which was fixed at 20 kHz, 400 W, the pulse duration was set to 2 s ON and 2 s OFF. The blank control sample was YPs suspensions with un-sonicated.

2.3. Particle size and zeta (ζ)-potential determination

For the particle size and the ζ-potential of YPs particles, following the method of Wang et al. [37] and Zhang et al. [38] with appropriate modifications. The particle size was measured using a laser scattering particle size analyzer (S3500, Microtrac Inc., Pennsylvania, USA) and Zetasizer Nano ZS90 (Malvern Instruments Co. Ltd., Malvern, UK). The average particle size was represented by D _[4,3]. The ζ-potential was measured at a room temperature of 25 °C using a Zetasizer Nano ZS90 (Malvern Instruments Co. Ltd., Malvern, UK). Before the measurement, the 5 % (w/v) YPs suspensions, including the sonicated samples were diluted to 0.2 % (w/v) with 10 mmol/L phosphate buffer (pH 7.0). Subsequently, the mixture was passed through a 0.45 μm Millipore membrane.

2.4. Turbidity determination

The turbidity of the YPs suspensions was determined based on the method of Gao, Rao, and Chen [39].The turbidity of each sample was assessed by measuring the absorbance at 600 nm using a UV-2600 spectrophotometer (Shimadzu, Japan). A blank solution consisting of ultrapure water was utilized.

2.5. Transmission electron microscope (TEM)

The YPs particles was observed with the transmission electron microscope (HT7800, Hitachi Ltd, Tokyo, Japan) operating at an acceleration voltage of 80 kV. The YPs suspensions in Section 2.2 was diluted 5-fold with 10 mmol/L phosphate buffer (pH 7.0). A copper grid was used to deposit 5 μL of YPs suspensions (1 % w/v), which were then stained with 2 % phosphotungstic acid for 120 s and dried at room temperature. They were subsequently used for the TEM observation.

2.6. Scanning electron microscope (SEM)

Surface morphology of the samples was obtained on a field-emission scanning electron microscope (SU-8010, Hitachi Ltd, Tokyo, Japan) at 5 kV. The YPs suspensions in Section 2.2 was lyophilized to yield the white powder. The YPs powder samples were attached to a double-sided carbon tape on a cylindrical aluminum mount, coated with a layer of gold.

2.7. Fourier-transform infrared (FT-IR) spectroscopy

FT-IR was achieved based on our previous method with few modifications [40]. The YPs powder samples are from section 2.6.

2.8. Intrinsic fluorescence spectra

The 5 % (w/v) YPs suspensions, including the sonicated samples were diluted to 0.2 % (w/v) with 10 mmol/L phosphate buffer (pH 7.0). Using a fluorescence spectrophotometer (RF-6000, Hitachi Ltd, Tokyo, Japan), the fluorescence emission spectra ranging from 200 to 450 nm were measured while keeping the excitation wavelength fixed at 280 nm. The width of both the excitation slit and the emission slit was set to 5 nm.

2.9. Surface hydrophobicity (H₀)

The ANS (1-anilino-8-naphthalenesulfonate) fluorescence probe was utilized to measure the surface hydrophobicity, following the approach outlined by Chen, Chen, Ren, and Zhao [41] with minor adjustments. The YPs suspensions were diluted to achieve various concentration (0.25 %, 0. 5 %, 0.75 %, 1 %and 1.25 % w/v) with 10 mM phosphate buffer at pH 7. Next, 20 μL solution of ANS with a concentration of 8 mM was prepared in the identical buffer and subsequently was introduced into the YPs suspensions, with a volume of 4 ml. The mixture was then thoroughly combined. The blends were stored in darkness for a duration of 15 min at ambient temperature. The extrinsic fluorescence intensity was measured using a fluorescence spectrophotometer (RF-6000, Hitachi Ltd, Tokyo, Japan) with an excitation wavelength of 390 nm and the emission spectrum was collected at 470 nm. The H₀ was determined by the slope of the regression curve between fluorescence intensity and protein concentration.

2.10. Interfacial tension

The pendant drop method was employed with the aid of a drop shape analyzer (DSA-100, KRÜSS GmbH, Hamburg, Germany) to determine the interfacial tension of YPs particles. Briefly, the needle of the syringe containing YPs suspension was first immersed in a quartz vessel contained grade 1 soybean oil. Then a drop of the YPs suspensions was created at the tip of a needle. The interfacial tension was obtained by quickly acquiring the drop image, detecting the edge, and fitting the Laplace-Young equation.

2.11. Wettability

The cylindrical tablets (10 mm × 2 mm) were formed by compressing the lyophilized YPs powder samples. The static sessile drop method was employed with the aid of a drop shape analyzer (DSA-100, KRÜSS GmbH, Hamburg, Germany) to measure the three-phase contact angle (θ) of particles. Fabricated tablets were positioned at the bottom of grade 1 soybean oil phase. The tiny water droplets (2 μL) were generated within the oil phase by the needle's tip and subsequently brought into contact with the tablet. After retracting the needle, a water droplet remained on the tablet. The image of the pendant drop was captured using a CCD camera that had a macro lens and subsequently examined with the DSA software.

2.12. Preparation of HIPPEs

HIPPEs were prepared with 5 % (w/v) un-sonicated and sonicated YPs suspensions (sonication time = 3, 7, 11, 15 min) (25 g) with grade 1 soybean oil (75 g) by passing through a homogenizer (IKA Ultra-Turrax T18, Staufen, Germany) at a speed of 15000 rpm for 2.5 min. These HIPPEs samples were named as control, S3, S7, S11 and S15, respectively. To prevent the growth of microorganisms and bacteria, a small amount of sodium azide (0.02 % w/v) was introduced into the mixture prior the homogeneity. The HIPEEs samples obtained were stored at 4 °C for further analysis.

2.13. Macroscopic morphology and microstructure observation

Macroscopic morphology: Macroscopic images were taken using vivo X60 smartphone (Vivo Mobile Communication Co., Ltd., Dongguan, China).

Confocal laser scanning microscope (CLSM) measurement: Confocal imaging was performed in a TCS Leica SP8 CLSM (Leica Microsystems, Wetzlar, Germany). The YPs and grade 1 soybean oil in freshly prepared HIPPEs were dyed separately using 10 μL of Nile blue and Nile red (1.0 mg/mL in isopropyl alcohol) for a duration of 30 min. 20 μL of stained HIPPEs was placed on slides and observed at 488 nm (Nile Red) and 633 nm (Nile blue), respectively.

Cryo-scanning electron microscopy (Cryo-SEM) measurement: The micromorphology of freshly prepared HIPPEs were evaluated by Cryo-SEM (SU-8010, Hitachi, Tokyo, Japan). Briefly, the HIPPEs samples in Section 2.12 were frozen by slicing in liquid nitrogen and subsequently the sublimated HIPPEs were coated with platinum before being placed inside a cold module chamber for observation.

2.14. Droplet size

To prevent agglomeration and flocculation of the droplets, the freshly prepared HIPPEs were diluted 20-fold with 1 % sodium dodecyl sulfate (SDS) solution, following a slight modification according to the method of Wu et al. [42]. The particle size was measured using a laser scattering particle size analyzer (S3500, Microtrac Inc., Pennsylvania, USA). The average droplet size was represented by D _[4,3]. D _[3,2] represented the diameter of HIPPEs droplets. The refractive indices of water and grade 1 soybean oil were 1.33 and 1.47, respectively.

2.15. Adsorption protein percentage (AP)

The freshly prepared HIPPEs were spun in a centrifuge (ST 16R, Thermo Scientific, MA, USA) at a speed of 10,000 rpm for a duration of 30 min. Following centrifugation, the Pickering emulsions exhibited the presence of oil, emulsion, and aqueous layers. Using a syringe, the transparent liquid layer at the bottom was taken and filtered through a 0.45 μm membrane. The protein concentration of the filtrate was determined using the Lowry method, with BSA serving as the standard. Then, the adsorbed proteins percentage [44]:

$$\mathit{AP}(\%)=\frac{C_0-C_1}{C_0}\times 100$$ (1)

where C_o is the total content of YPs proteins in the where before centrifugation, C₁ is the content of YPs proteins in the aqueous phase after centrifugation.

2.16. Rheological behavior

The rotational rheometer (MARS60, HAAKE, Germany) was utilized to examine the rheological characteristics of HIPPEs, following the approach outlined by Zhang et al., [45]. A moderate amount of the HIPPEs sample was positioned between two parallel plates, and the apparent viscosity was assessed within the range of shear rates from 0.1 to100 s⁻¹. The storage modulus (G′) and loss modulus (G″) were obtained by frequency sweeps (0.1–13 Hz frequency and 1 % strain).

2.17. Statistical analysis

Measurements were conducted in triplicate, which is three independent measurements from the same sample, and the outcomes were reported as the average plus the standard deviation (SD). The IBM SPSS 20.0 Statistical Software Program (SPSS, Inc., Chicago, IL, USA) was used to conduct one-way an analysis of variance (ANOVA) on the data with Duncan’s test (P < 0.05). The figures were created using Origin 2018 software from Microcal, a company based in the United States.

3. Results and discussion

3.1. Formation and characterization of YPs particles

3.1.1. Particle size

Particle size and distribution are critical indicators to reflect the success of YPs particles prepared by the sonication [47]. The size and distribution of the protein particles is closely related to the stability of HIPPEs, due to it affects the adsorption and arrangement of protein particles at oil/water interface [35], [49]. Fig. 1A-B and Fig. 1C respectively depicted the effect of ultrasonic time on particle size distribution and average particle size of YPs. From Fig. 1A and Fig. 1B, two particle size distribution peaks of YPs were both blue-shifted compared with the un-sonicated sample after sonicated for 3–11 min at 400 W. The particle size of YPs particles of sonication markedly decreased compared to the un-sonicated sample as indicated in Fig. 1C. This decrease and blue-shifted in particle size of YPs particles may be attributed to the shear forces, shock waves, and microflows generated during by the mechanical vibration of ultrasound [50], [51]. They are able to open up the tightly-packed structure of YPs aggregates in the aqueous solution through the disruption of the inter- or intra-molecular non-covalent interactions between YPs molecules, such as the hydrogen bond, hydrophobic interaction and electrostatic interaction [52], [53]. This enhanced the stability of YPs, because it may impact the mobility of the particles in the continuous phase as well as the droplet size, rheological properties and stability of HIPPEs [55]. Meanwhile, their average particle size first decreased rapidly and then increased with an increase in sonication time, and the minimum particle size was obtained as sonicated for 7 min. Compared to sonicated for 7 min, too long of a sonication could lead to the formation of larger YPs particles due to increase the rate of Brownian movement and collision [48], [56]. Alternatively, too long of a sonication increased the chances of collision between YPs molecules or small particles to form an aggregate [57]. Physically, these mean that the aggregation rate greater than fragmentation rate for YPs particles in the solution [58]. This aggregation was primarily driven by electrostatic and hydrophobic interactions [59]. Additionally, several studies argued that the longer ultrasonication time results in heat-induced aggregation of small particles, which has been attributed to the creation of high temperature [60], [61]. However, sonication was performed in an ice bath in this experiment, and hence YPs particles were not occurred heat-induced aggregation.

Fig. 1.

Particle size distribution (A, B), average particle size (D _[4,3]) (C) and turbidity (D) of YPs particles prepared at different sonication time. Note: Different letters on the bars indicate significant differences (P < 0.05).

3.1.2. Turbidity and microscopic morphology

Turbidity is a macroscopic parameter that measures the level of protein aggregation, which mirrored the aggregation state of the YPs particles in aqueous solution to some extent [14]. Generally, higher turbidity corresponds to denser protein particles aggregation, and it is negatively correlated with stability of high internal phase Pickering emulsion prepared from protein particles [62]. TEM was used to visualize the microscopic morphology of YPs in aqueous solution after sonication. The effects of different sonication time on the turbidity of YPs aqueous solution were shown in Fig. 1D. From TEM images (Fig. 2A), as expected, it was that YPs in all aqueous solution samples exist in two distinct states: small globular (top) and large bulk (bottom). In particular, compared with the microstructure of the un-sonicated sample, ultrasound transformed the large bulk YPs into a flocculent in aqueous solution. The turbidity was significantly decreased in all sonicated samples (P < 0.05). Which shows that ultrasound was capable of altering the aggregation morphology and level of YPs, and they were related to the sonication time. The cavitation effect generated by ultrasound reduced the size of YPs particles (Fig. 1C). Leading to the specific surface area available for light scattering increased, thus decreasing the turbidity of YPs aqueous solution [63]. The size of small particles and floccules turned to be smaller first and then larger with progressive prolongs of sonication time. And turbidity decreased initially and then increased. Obviously, compared to other sonication time, 7 min was more conducive to the formation of small YPs particles/flocculent, coinciding with the results of particle size distribution above. Owing to the strong physical force of the ultrasonic waves, the inter- and intra-molecular interacting force (such as hydrogen bonding, hydrophobic interactions, electrostatic interactions, and van der Waals forces) may be disrupted [64]. Which resulted in a decreased size of YPs aggregates and a decrease in turbidity. However, prolonged sonication (15 min), the probability of the particles collided and rubbed with each other substantial increased. Resulting a faster aggregation rate of YPs molecules than the decomposition rate of YPs flocculent, the turbidity increases terminally. To further verify the formation of YPs particles, the microstructure and morphology of YPs lyophilized powders displayed by SEM. The resulting micrographs (Fig. 2B) clearly showed the YPs lyophilized powders were composed of two populations of particles, namely large spherical particle and small spherical particle. Meanwhile, their size first decreased and then increased with increasing sonication time. Thus, the effect of sonication time on particle preparation of YPs were further confirmed by microstructure results.

Fig. 2.

TEM (A) and SEM (B) images of YPs particles prepared at different sonication time.

3.1.3. ζ-potential analysis

The ζ-potential was employed to indicate the net surface charge of YPs, the higher absolute ζ-potential, the stronger repulsive force between molecules, which is one of the most important factors in determining dispersion and aggregation of YPs [65]. As shown in Fig. 3, the un-sonicated sample had a negative ζ-potential at pH 7.0. Upon ultrasonic treatment, the absolute ζ-potential value of the YPs firstly increased and then went through a slight decrease with increased sonication time with the turning-point occurred under the sonication time of 7 min. Suggesting that negatively charged amino acid residues on YPs surface gradually increased and then progressively decrease. The possible explanation for this result is that sonication unfolded the YPs structure, such as hydrogen bonding between polar amino acid residues and hydrophobic interactions between non-polar residues are disrupted [39], [54]. This contributes to increasing negative surface charge resulted in an increase in the absolute ζ-potential value. The prolonged sonication caused the aggregation of yeast protein particles. Consequently, a portion of negatively charged amino acid residues of YPs surface were buried, the surface negative charges decrease at 15 min of sonication and the absolute ζ-potential value decrease [66]. It has previously been reported that a proteins dispersion system with a higher ζ-potential is electrically stabilized while a lower ζ-potential tends to promote aggregation [67]. This was consistent with the result of the turbidity measurement and TEM of this study. Nevertheless, the effects of sonication time on conformation of YPs, intensive investigations are still needed. Since the molecular structural changes and the intra- and intermolecular interactions of YPs could have consequences for the aggregation behavior of YPs particles at the HIPPEs interface [68].

Fig. 3.

ζ-potential of YPs particles prepared at different sonication time. Note: Different letters on the bars indicate significant differences (P < 0.05).

3.2. Structural properties of YPs

3.2.1. Intrinsic fluorescence spectroscopy

The tertiary conformation changes of protein were characterized by measuring their intrinsic fluorescence, which is yielded from the emission of the three amino acids with aromatic ring side chains—phenylalanine (Phe), tyrosine (Tyr), and especially tryptophan (Trp) fluorescence spectrum [69]. The maximum fluorescence intensity (I_max) and maximum emission wavelength (λ_max) can evince the changes of fluorescence quantum yield of chromophores (Phe, Tyr and Trp) with varied environmental hydrophobicity [70], [71]. Normally, chromophores are in a hydrophilic environment if λ_max is greater than 330 nm [72]. The effects of different sonication time on the intrinsic fluorescence spectra of YPs molecules were shown in Fig. 4. The I_max increased and then decreased drastically with duration of sonication time when compared to un-sonicated sample. The highest I_max appeared in the sample of sonication 11 min. As same time, the λ_max treated with ultrasound first displayed a slight blueshift and then returned to the original wavelength could be noticed. The increased I_max may be due to increase the emission through the sonication enhanced exposure of the aromatic amino acid residues in the sample [73]. Afterwards, because extra input of ultrasonic energy added the probability of collision and assembling of protein molecules, provoking YPs aggregation [74]. The largest blue shift of λ_max was 1, 1, and 0.5 nm in YPs suspensions under the ultrasound time of 3 min, 7 min, and 15 min, respectively (Fig. 4). The blue shift of λ_max means that more aromatic amino acid residues are exposed to the internal relatively hydrophobic environment [75]. It is normally indicating aggregation of proteins, while the red shift indicates the occurrence of depolymerization phenomena of the protein aggregates [60]. Which was contrasts the particle size distribution and the microscopic morphology presented in this study. Hypothesizing that this may be due to the relatively few number of aromatic amino acid residues in YPs. Therefore, an extrinsic fluorescence probe (ANS) that binds nonspecifically to the hydrophobic areas of protein molecule exposed the aqueous solvent was utilized to measure yeast protein surface hydrophobicity (H₀).

Fig. 4.

Intrinsic fluorescence of YPs prepared at different sonication time.

3.2.2. Surface hydrophobicity (H₀)

The influences of the sonication on the H₀ of YPs are displayed in Fig. 5. Upon sonication, an increase in the H₀ of the YPs followed by a final decrease compared with the un-sonicated sample. The H₀ of the YPs with sonication for 7 min was the highest. The increase in H₀ was attributed to the promotion of the breakdown of the macromolecular particles of YPs upon cavitation and acoustic streaming, thereby leading to the unfolding of protein molecular structure [76]. Subsequently, the hydrophobic regions and groups to be more exposed on the surface of protein molecule, thereby improving the hydrophobic capacity of YPs [77], [78]. The H₀ of samples treated for 11 and 15 min reduced dramatically compared to samples sonicated for 7 min. This was attributed to the aggregation of the small YPs or unfolded YPs molecules caused by over sonication, which mainly through intermolecular attractive hydrophobic interactions, burying the hydrophobic group [70], [79]. Notably, the trend of changes in the H₀ of yeast protein was basically corresponded to the I_max and contrary to the λ_max. A reasonable explanation for this inconsistence might be that the decomposition of large YPs into small YPs leads to an increase in surface area. This was potentially offering additional binding sites for ANS probe, resulting in higher values of H₀ [80]. Thus, the combination of turbulence, shear forces, high pressure, and shock waves effects by the sonication produces had an impact on the tertiary and quaternary structures of YPs.

Fig. 5.

Surface hydrophobicity of YPs prepared at different sonication time. Note: Different letters on the bars indicate significant differences (P < 0.05).

3.2.3. FTIR spectroscopy

FTIR spectroscopy is an effective means to determine structure conformations by detecting the vibrational spectra of the local environment of YPs side chains [13]. At the same time, it could quantify the secondary structural fractions changes of YPs molecules specifically [82]. Secondary structure analysis by FTIR spectroscopy (ranging between 4000 and 500 cm⁻¹) confirmed the effect of sonication on the structural properties of YPs (Fig. 6A). Various distinct bands in the infrared spectrum represent various highly polar bonds or functional groups in the YPs. The main spectral observations were four intense bands attributed to amide A (3400–3200 cm⁻¹), amide B (2980–2850 cm⁻¹), amide I (1700– 1600 cm⁻¹), and amide Ⅱ (1550–1450 cm⁻¹) [83], [84]. In general, the broad peak between 3200 and 3600 cm⁻¹ relates to the inter- and intra-molecular hydrogen bonds. They represent the stretching vibration of O–H and N–H bands of amide A in YPs molecules as well as in YPs and water molecule interaction. The shift of the amide A indicated that O–H bonds and N–H bonds were involved in the reaction [85]. And it indicated that sonication influenced the intra- and intermolecular interactions of YPs and corresponding changes of protein structure. The amide A of YPs with sonication was blue shifted compared to the un-sonicated sample. Meanwhile, compared with the un-sonicated sample, the intensity of the peak at 3200–3600 cm⁻¹ of the sample with sonication for 7 min slightly increased, while the intensity of peak of the other samples with sonication decreased. This suggested that the presence of more hydrogen bonds in the sample with sonication for 7 min, which was more hydrophilic than the other samples [86], [123]. The amide B at 2980–2850 cm⁻¹ was attributed to stretching vibrations of C–H in –CH₃ and –CH₂ of the YPs molecules [19], [20]. The amide Ⅱ was attributed to the bending vibration of N–H and the stretching vibration of C–N. The absorption of the amide I (1700–1600 cm⁻¹) is due to stretching vibrations of C = O and C–N in the YPs structure [19], [87]. It represents the aggregation, folding, and unfolding of the YPs sonicated and using to extract information on YPs secondary structures [88].

Fig. 6.

FTIR spectroscopy of YPs prepared at different sonication time. Note: Different letters on the bars indicate significant differences (P < 0.05).

The amide I was deconvoluted by the second derivative method to quantify the contents of the secondary structure of each sample of YPs. The percentages of ɑ-helix, β-sheet, β-turn, and random coil conformation of samples were demonstrated in Fig. 6B. The results showed that the β-sheet and β-turn were the abundant secondary structure element of the YPs, followed by α-helix and random coil. After sonication, ɑ-helix and random coil content of YPs was found to first decreased and then increased with increase in sonication time, an opposite tendency in β-sheet and β-turn content was observed. The ultrasonic cavitation generated intensive hydrodynamic shear in the local area, which could lead to interconvert between secondary structural fractions of YPs, dissociation of the protein conformation, and remodeling of the proteins [10]. However, in comparison to the un-sonicated sample, the trend of the secondary structure fractions content in YPs with sonication for 7 min was significantly different from that of other sonication groups. Specifically, the β-sheet content was found to significantly decreased for YPs with sonication for 7 min compared with the un-sonicated sample, β-turn content was significantly higher, and no significant difference was observed for α-helix and random coil content. This attributed to the ultrasound disrupted the hydrogen bond that mainly maintains the β-sheet conformation in protein, causing a decrease in the β-sheet content [81], [82]. This also leads to the YPs conformation was dissociated and tends to unfold, exposing its glycine or proline residue, resulting in the formation of β-turn [89], [90]. Their exposure is also one of the reasons for the increased in H₀ of YPs. Furthermore, from Fig. 6B, the sum of α-helix content and β-sheet content of YPs with sonication for 7 min was lower than other sonication groups and the un-sonicated sample. The conformational stability of YPs is influenced by α-helix and β-sheet, which are rigid structures found within the polypeptide chains. On the other hand, the flexibility of YPs was attributed to β-turn and random coil [91]. Cheng et al. [3] argued that the increase of flexible of protein implies unfolding of protein and is usually beneficial to the improvement of the protein functional properties. Therefore, it was speculated sonication for 7 min could induce unfolding and flexible conformational reorganization of the YPs, thereby exhibiting better flexible properties. This might be more beneficial to the adsorption behavior of YPs particles on the oil/water interface of HIPPEs.

3.3. Interface property of YPs particles

3.3.1. Interfacial wettability

Similar to the amphiphilicity of surfactants, the wettability of YPs particles plays a vital role in assessing the stability of HIPPEs [92], [55]. The three-phase contact angle (θ) is main parameters characterizing wettability [27]. When the θ is between 15° and 90°, the YPs particles exhibit a hydrophilic characteristic. Conversely, when the θ falls between 90° and 165°, the particles possess a hydrophobic nature according to Dickinson [93]. Ideally, YPs particles possessing an angle of approximately 90° exhibit a nearly neutral wetness at the oil/water interface, which is more suitable for creating stable HIPPEs [94]. The results are shown in Fig. 7, the θ of un-sonicated yeast protein particles was 98.17°, indicating that the predominance of YPs particles hydrophobicity. Compared with that of the un-sonicated yeast protein particles, the θ of sonicated YPs particles were decreased by 9.26–27.22°, suggesting that the hydrophilicity of YPs particles with sonication was effectively improved. However, the results of H₀ showed that the hydrophobicity of YPs particles also increased after sonication. This illustrates that the changes in the hydrophobic-hydrophilic character of YPs particles after sonication of YPs might lead to assume new equilibrium structures. Specifically, the application of sonication induced the YPs structure to unfold and exposed hydrophobic amino acid residues and more polar groups buried within the YPs to some extent, leading to a decrease in the θ [73]. Moreover, the θ of YPs particles presents a trend of increase first and then decrease with the sonication time prolonged. When the sonication time was increased from 3 min to 7 min, the θ increased from 73.57° to 88.91°. From TEM (Fig. 2A) and SEM (Fig. 2B) images, this was due to sonication at this stage reduced the level of YPs aggregation, which lead to a more unfolded protein structure. This may promoted the release of hydrophobic amino acids residues higher than hydrophilic amino acid residues at this stage [43]. The H₀ results support this trend. But ultimately, the amount of hydrophobic amino acid residues in YPs sonicated 7 min was still slightly lower than that of hydrophilic amino acid residues. The change of hydrogen bonds in amide A band could demonstrate that YPs had the ability to bind to water when subjected to sonication for 7 min. Hence, its θ close to 90° suggests that the wettability is close to neutral. In this situation, it might contribute to the firmer adsorption of YPs particles at the interface and the formation of a spatial barrier, preventing droplet aggregation in an emulsion [95]. With the increase in sonication time (>7 min), the θ of YPs particles showed a decreasing trend. Combining TEM and H₀ results, this may be attributed to recombination of YPs molecules and aggregation of small YPs particles caused by prolonged sonication. Some hydrophobic groups were buried, exposing more hydrophilic groups on the surface of YPs particles, which led to a decrease in the θ [96]. Overall, the sonication improved the wettability of YPs particles, and the sample subjected to 7 min of sonication had near-perfect amphiphilicity.

Fig. 7.

The three-phase contact angle of YPs particles prepared at different sonication time. Note: Different letters on a graph indicate significant differences (P < 0.05).

3.3.2. Interfacial tension

Interfacial tension can serve as an indicator to characterize the surface properties of YPs particles and interfacial interactions, because it elucidates the interfacial adsorption behavior of protein particles [97]. It plays a crucial role in the stability of HIPPEs. As shown in Fig. 8, the trend for changes in interfacial tension of YPs were divided in two different stages that include the interfacial tension drops (initial period) and the interfacial tension tended to a plateau (second stage). The decrease in interfacial tension of each sample in the initial period was caused by the adsorption and aggregation processes of YPs particles on the bare interface [98]. The interfacial tension tended to a plateau was due to the steric hindrance and electrostatic energy generated by the adsorbed YPs particles, which impeded the subsequent adsorption [99]. Experimental observations, compared with the YPs particles with sonicated, indicate that the time needed for the interface tension of the un-sonicated YPs particles to reach the second stage was longer. Base on the reasons given above, this was created by the θ of 98.17° of the un-sonicated sample and the lower ζ- potential absolute value. Again, it also points to the sonication may enhance the absorption, stretching, and reorganization of YPs molecules at the oil/water interface. Notably, in the sonicated group, the more flexible and expanded structure conformation of YPs with sonication for 7 min, and it displayed smaller particle size and near-neutral wettability. This was beneficial for the adsorption and ordered arrangement of YPs at the oil/water interface, reducing the interface free energy and improving the interface stability [99]. However, the results of the sample of sonicated for 15 min are the opposite. Therefore, compared with other YPs samples with sonicated, the interfacial tension value of the YPs with sonication for 7 min was lower. The results summarized in this section demonstrate that the YPs particles prepared through an appropriate time of sonication may contribute to the formation of stable HIPPEs.

Fig. 8.

The interfacial tension of YPs particles prepared at different sonication time.

3.4. Characterization of HIPPEs

3.4.1. Hippes formation

In this section, the HIPPEs was prepared using YPs particles as stabilizer and soybean oil as the internal phase. The macroscopic stability of HIPPEs was first characterized by observing its macroscopic morphology. At the macroscopic level, all YPs-stabilized HIPPEs showed a relatively stable steady state due to there was no flow behavior observed when they were inverted (Fig. 9A). CLSM enabled the direct observation of the YPs network structure within the HIPPEs and the morphology of oil droplets [100]. Additionally, the capability of CLSM to observe the adsorption and positioning of protein particles at the oil/water interface of Pickering emulsions has been proven [101]. The difference in the microstructures of HIPPEs stabilized by YPs particles (prepared at different sonication time) was detected using CLSM. The result was presented in Fig. 9B. The red indicated the existence of internal oil phase within the HIPPEs droplets, while the green represented the YPs particles that covered the surface of the HIPPEs droplets. A merge of the above two images was shown with another color. As can be seen that in all samples, the spherical or polyhedral morphology and closely packed of the oil droplets within the protein network, and it confirmed the formation of O/W HIPPEs. From their enlarged images, it could be seen that there was a YPs particles wrapping layer (green) around the oil droplet, which was connected to the YPs particles gathered outside. The boundary of the wrapping layer was relatively clear. However, the amount of YPs particles that can be adsorbed to the oil/water interface is limited, and the remaining particles remain in the continuous phase as non-adsorbed particles. These non-adsorbed particles accumulating and acting as network structures to stabilize HIPPEs droplets [102]. Obviously, in comparison to control, the oil phase distribution was more dispersed and the sizes of the oil droplets became smaller in S3 and S7 samples, especially for the S7, as observed in the red images. The oil droplet size was increased gradually and more closely arranged in S11 and S15 samples. It shows that the thickness of interfacial layer formed by yeast protein particles at the oil/water interface was increase initially and then decrease with increased sonication time. The same phenomena can also be observed from the corresponding green images. It was primarily caused by differences in the particle size, wettability, interfacial tension, and H₀ of different YPs particles [103]. However, this difference was attributed to changes in the molecular structure of these YPs caused by different sonication time, such as intrinsic fluorescence spectroscopy and FTIR spectroscopy. Normally, small particle sizes and near-neutral wettability were more beneficial to the adsorption of YPs particles to the oil–water interface and fully unfold at oil–water interface [104]. Concomitantly, they rely on hydrophobic interaction to form complete and tight interfacial layers around the oil phase ([94]. Hence, S7 sample could obtained smaller droplet size and thicker interfacial layer. Notably, the oil droplet size of S15 sample was significantly larger compared to other samples, and its YPs showed obvious aggregation in the pores between the droplets. It was surmised that this may result from the coalescence of emulsion droplets. As a consequence, the stability mechanism of YPs particles for HIPPEs in this study was the bridged network mechanism as described by Ming et al. [105]. The YPs are closely held together by hydrophobic interaction to form interfacial protein layers or acted as the protein network. The coalescence stability of these HIPPEs primarily relies on the steric hindrance effect, preventing coalescence and aggregation between emulsion droplets [106].

Fig. 9.

Appearance photograph and CLSM images of the HIPPEs stabilized by YPs particles that were prepared at different sonication time.

3.4.2. Adsorption protein percentage (AP)

The thickness of the interface layer formed by YPs particles is related to the ease with which the coalescence of emulsion droplets. The AP is indirect means of reflecting the thickness of interfacial protein layer formed by YPs particles, it is negatively correlated with the coalescence probability of the HIPPEs [107]. The AP of HIPPEs was measured to verify the adsorption behavior in the YPs particles interface at different sonication time, as shown in Fig. 10. Compared with the control, appropriate sonication time (≤11 min) could significantly enhance the adsorbed protein percentage. This might be ascribed to the unfolding of the YPs conformation after sonication with appropriate time (Section 3.2.2), which could increase the flexibility and H₀ of YPs and promote the formation of YPs layer on the oil/water interface [108]. The S7 sample showed the highest AP. It can be attributed to the smaller size (Fig. 1C) and better particle wettability (Fig. 7) of YPs particles sonicated for 7 min. Due to their larger specific surface area, the smaller YPs particles could offer superior surface coverage when compared to particles of the same mass [109]. This is beneficial for the absorption of the YPs, resulting in a rise in the AP. Thus, the YPs particles formed an interfacial layer with higher mechanical strength on the oil/water interface to, which could withstand the pressing force between two approaching emulsion droplets, preventing the coalescence of the HIPPEs droplets [46]. However, at sonication time higher than 7 min, the AP of HIPPEs ultimately declines with increasing sonication time. The AP of S15 sample was even significantly less than control. This may be due to too long of a sonication conferring YPs particles a more compact conformation, larger particle size, lower H₀, as well as higher hydrophilicity and higher interfacial tension. These factors were unfavorable for the adsorption of YPs particles on the oil/water interface, leading to an increased probability of oil droplet coalescence and the micro stability of HIPPEs may deteriorate [110]. Therefore, an assessment of the emulsion droplets coalescence appeared essential for HIPPEs.

Fig. 10.

The adsorbed YPs percentage of the different HIPPEs. Note: Different letters on the bars indicate significant differences (P < 0.05).

3.4.3. Cut surface of HIPPEs

The cut surface structure each HIPPEs sample was further observed using Cryo-SEM, as shown in Fig. 11. When the YPs sonicated within 7 min (including 7 min), two or more oil droplets in HIPPEs sharing the same YPs particles interface can be clearly observed in control, S3 and S7 sample, which is an important feature of bridged network mechanism. As expected, a thick interfacial layer was observed for its stabilized HIPPEs when the YPs was sonicated 7 min. This is in excellent agreement with the AP results. However, the droplets of S11 and S15 sample showed a loose structure, and the spaces between the droplets were filled by the more honeycomb structure. This is attributed to the elevated concentrations of non-adsorbed YPs particles in the continuous phase (Section 3.4.2), they trend to creating a 3D network through a variety of intermolecular and interfacial interactions in the continuous phase to stabilize HIPPEs [36]. This transition of the stabilization mechanism is likely caused by the coalescence of HIPPEs droplets, because the interfacial layer formed by large YPs particles was unstable. The Cryo-SEM images showed good agreement with the CLSM images. Altogether, the findings from observing HIPPEs cut surface indicated that the YPs particles functioned as efficient food-grade Pickering stabilizers. However, the YPs particles, which were produced with varying sonication time, displayed distinct adsorption patterns and stabilization mechanisms.

Fig. 11.

The Cryo-SEM images of the different HIPPEs.

3.4.4. Droplet size distribution of HIPPEs

Droplet size and distribution is a crucial parameter when assessing HIPPEs systems, because it can reflect the degree of coalescence of emulsion [111]. Fig. 12A and Fig. 12B respectively showed the impact of ultrasonic time on the distribution and size of HIPPEs droplets. The data revealed the droplet size distribution peak of HIPPEs samples first shifted to the left and the mean droplet size (D _[4,3]) first decrease sharply with the extension of sonication time. The variation trend showed inflection points at 7 min of sonication. The droplet size distribution peak of HIPPEs samples immediately starts to shift to the right after YPs were sonicated exceeds 7 min, and the mean droplet size increases. This is an interesting phenomenon in this study, clearly, the smaller particle size of YPs (Fig. 1C) the smaller was the droplet size of the stabilized HIPPEs. This can be explained by the following two factors. One important point is that smaller particles exhibit a larger diffusion coefficient and rate of interface coverage, which makes them more advantageous for adsorption at the interface [55]. Thus, they could quickly form stronger interfaces through intermolecular interactions. The formation of smaller droplet size was facilitated. Another reason was that a large particle obviously cannot accommodate the large curvature of a small droplet, which results in they cannot form stable small droplets [112]. Despite the huge adsorption energy of larger YPs particles [113]. Additionally, the presence of steric hindrance among smaller YPs particles within distinct droplets also contributes to the deceleration of coalescence [35]. The small droplets are more beneficial to the stability of YPs particles-based HIPPEs. The likelihood of coalescence between neighboring droplets decreases [114]. Thus, the higher D _[4,3] values of S15 and S11 compared to other samples may be due to their coalescence.

Fig. 12.

Droplet size distribution (A) and average droplet size (D _[4,3]) (B) of the different HIPPEs. Note: Different letters on the bars indicate significant differences (P < 0.05).

3.5. Rheological behaviors of the HIPPEs

The rheological properties are regarded as a significant benchmark for evaluating HIPPEs appearance, performance and stability, which holds great significance in their utilization in the food industry [55]. Therefore, the apparent viscosity and viscoelastic properties of all the prepared HIPPEs were investigated by shear rate sweep test and frequency sweep test. The viscosity properties of HIPPEs have always been a vital basis in analyzing the droplet–droplet coalescence. The apparent viscosity as a function of shear rate was shown in Fig. 13A. All HIPPEs samples exhibited the typical non-Newtonian pseudoplastic fluid and shear thinning behavior as the variation of shear rates from 0.1 to 100 s⁻¹. The shear-thinning behavior was an indicator of the gradual disruption of the HIPPEs droplets [115]. These results reflected the deformation of the oil droplets and the breakdown or rearrangement of internal structure of the HIPPEs during the shearing process [116]. The apparent viscosity of HIPPEs stabilized by the YPs particles was affected by the sonication time used to prepare YPs particles. It was observed that S7 exhibited slightly high apparent viscosity as compared to that of control, while S15 exhibited a slightly low apparent viscosity. This phenomenon was attributed mostly to the difference in these HIPPEs structure. Specifically, an increased in the AP value of the sample, and the formation of crosslinking or bridges between droplets (Cryo-SEM images) both would increase the apparent viscosity of HIPPEs [117]. And reciprocally, the apparent viscosity of the HIPPEs decreased.

Fig. 13.

Apparent viscosity (A) and Frequency sweep (B) of the different HIPPEs.

Apart from the viscosity characteristic, the storage (G′) and loss modulus (G″) are equally crucial to assess the HIPPEs stability and quality. The influence of the YPs particles, treated at different ultrasonic time, on the viscoelasticity of HIPPEs were investigated using dynamic oscillatory measurements. The dynamic oscillatory data (Fig. 13B) showed that the G′ of all samples was higher than the G″, confirming the elasticity-dominated attributes or solid-like properties of HIPPEs stabilized by the YPs particles [118]. For all samples, the G′ and G″ values were disjoint and both increased with a gradual increase in angular frequency. The S7 exhibited comparatively high G′ than other samples, while S15 exhibited a relatively low G′ than other samples. Simultaneously, no discernable difference was seen in G′ in the S3 and S11 compared to control. There was little difference in the G″ from each sample, the S7 slightly higher than those found at other samples. The elasticity of HIPPEs is typically associated with the interfacial characteristics, the particle network within the continuous phase, and the droplet size [119]. Given the preceding results (Section 3.4), the S7 sample possessed the smaller droplet size, a thick interfacial protein layer, and an even 3D network structure. This resulted in the lower droplet fluidity and increased resistance to deformation and ultimately improve G′ values of S7 sample [10]. Moreover, the S15 showed a loose structure due to droplet coalescence (Section 3.4.3), which caused a dramatic reduction in resistance to deformation, resulting in lower G′. Therefore, the rheological results suggested that the sonication time played an important role in the rheological properties and stability of HIPPEs stabilized by the YPs particles. The best outcome was achieved with the sonication on YPs for 7 min to stable HIPPEs in this experiment.

3.6. Coalescence mechanism of HIPPEs

In general, the droplet coalescence of HIPPEs can be divided into two consecutive processes: packing and fusion [120]. The droplets of HIPPEs press against each other due to higher volume fraction of the internal phase, leading to transform them from spheres to rounded polyhedrons. In order to achieve the close packing of droplets for coalescence, however, Laplace pressure (P₀) must be overcome by the external pressure provided by the droplets’ weight (P), as illustrated in Eq. (2) [120]. The fusion occurs when the interfacial particle layer ruptures, causing the droplets to merge. In this Section, therefore, the coalescence stability of each HIPPEs sample was evaluated in terms of Eq. (2).

$$P>P0;P0=\frac{4\gamma }{d}$$ (2)

where γ is interfacial tension of Pickering particles at the oil/water interface, d is the droplet diameter (D _[3,2]) of Pickering emulsion.

From Eq. (2), P₀ decreases with increasing droplet size of emulsion. However, it is well known that the P increases with increasing droplet diameter of emulsion. In other words, when the interfacial tension of YPs particles remains constant, the smaller the droplet size, the lower the probability of HIPPEs coalescing. Conversely, the higher the probability of HIPPES coalescing. Evidence for this notion was present in the literature ([121], [122]. The D _[3,2] values of each HIPPEs sample are shown in Table 1. The D _[3,2] value of HIPPEs first decrease and then increases in response to the increasing with sonication time. This was mainly determined by the differences in the structure, size, ζ-potential, wettability, and absorption protein percentage of YPs particles caused by different sonication time. Moreover, in spite of the YPs particles with sonication for 11 and 15 min showed the higher interface tension value compared with those of the sonication for 7 min, their differences were not obvious (Fig. 9). However, S11 and S15 sample showed a 2-fold droplet size value over S7 sample (Fig. 13B). Therefore, according to Eq. (2), the coalescence probability of droplet in S15 and S11 was much higher than that in the control, S3 and S7. This confirmed previous observations and speculations that the YPs particles in the control, S3 and S7 primarily form a close-packed interfacial layer through adsorption to stabilize HIPPEs and against droplet coalescence. But the stability mechanism of YPs particles in S11 and S15 appears to be altered compared to control, S3 and S7. The network framework which is composed of non-adsorbed YPs particles in the continuous phase appears to play a dominant role in the stability mechanism, rather than interfacial protein layer.

Table 1.

The droplet diameter (D _[3,2]) of the HIPPEs stabilized by YPs particles that were prepared at different sonication time.

	HIPPEs samples
	control	S3	S7	S11	S15
D [3,2] (μm)	32.33 ± 1.52a	24.15 ± 0.99c	21.48 ± 0.82d	29.90 ± 1.66b	34.35 ± 0.95a

Note: Different letters indicate significant differences (P < 0.05).

4. Conclusion

Taken together, the sonication was able to modulate the molecular structure and conformation of YPs, thereby improving the physical and chemical properties of commercial YPs as Pickering particles. Including mainly particle size, surface charge, surface hydrophobicity, surface wettability, interfacial tension and so on. The results indicated that YPs sonicated for 7 min exhibited a more relaxed molecular structures and conformation, smallest particle size, and optimal amphiphilicity. Mechanistically, it was revealed that the YPs particles mainly stabilize HIPPEs through bridging mechanisms. The YPs particles adsorbed on the oil/water interface spontaneously form an interfacial protein layer through hydrophobic interactions. The non-adsorbed YPs particles occurred aggregation in the continuous phase and act as “structural agents” in bridging systems. Forming a 3D protein network framework with the interface protein layer through hydrophobic interactions, electrostatic interactions, or steric hindrance effects to stabilize oil droplets in HIPPEs. The HIPPEs stabilized by YPs particles sonicated for 7 min exhibited the highest adsorption interface protein percentage and a more homogeneous 3D protein network, resulting in the smallest droplet size and the highest storage (G′). The transition from extended to compact conformations of YPs occurred when the sonication time exceeded 7 min. Ultimately leading to a decrease in the amount of adsorbed protein at the oil/water interface. The droplets of HIPPEs stabilized by them were prone to droplet coalescence and the droplet diameter increases, but no demulsification phenomenon. The non-adsorbed YPs particles form larger aggregation structures in the continuous phase. In all, for the stabilization mechanism of HIPPEs, the contribution of the interface protein layer formed by yeast protein particles sonicated for 7 min in bridging system was relatively greater. Whereas “structural agent” contributed more to bridging system when the sonication time exceeded 7 min.

CRediT authorship contribution statement

Tianfu Cheng: Writing – original draft, Software, Conceptualization. Guofang Zhang: Visualization, Validation, Supervision. Fuwei Sun: Data curation. Yanan Guo: Methodology. Ramnarain Ramakrishna: Writing – review & editing. Linyi Zhou: Investigation. Zengwang Guo: Project administration, Funding acquisition. Zhongjiang Wang: Resources, Funding acquisition.

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.