Abstract
Protein-based nanoparticles have gained attention in stabilising emulsions since it has a wide range of application. This study utilised rice bran protein (RBP) as a raw material to form RBP fibrils via acid heat treatment. Transmission electron microscopy revealed after 6 hr of heating, RBP adopted an elongated fibrillar structure. During fibril formation, an increase in thioflavin T fluorescence intensity was observed, along with the emergence of numerous well-organised β-sheet configurations. The water-in-oil-in-water (W/O/W) double emulsions prepared during the fibrils formation were analysed by particle size, viscosity, and laser coaggregation microscopy. The results indicate that, compared to unheated RBP, emulsions prepared with RBP fibers exhibited reduced aggregation, a significantly smaller particle size, significantly increased viscosity, a greater thickness of RBP fibers at the oil–water interface, and significantly enhanced stability (p < .05). The improved emulsion characteristics were directly linked to the interfacial features of the RBP fibrils. This study provides a method for constructing stable W/O/W ratios.
Introduction
Rice bran is an organic food item that is obtained as a by-product of the rice milling process. It contains many healthy substances, such as proteins (14%–16%) (Zhang et al., 2022), carbohydrates (33%–42%), and fats (12%–23%), and the composition of the substance includes a significant amount of dietary fibrils (25–40%) and phenolics (9.60–81.85 mg GAE/g; Punia et al., 2021). Therefore, various functional components can be extracted from rice bran using physical extraction methods, chemical extraction methods, biological extraction methods, as well as the rapidly developing green extraction technologies in recent years, such as ultrasonic extraction, supercritical fluid extraction, and enzymatic hydrolysis (Garza-Cadena et al., 2023). These components can be utilised for the preparation and development of functional foods or cosmetic formulations (Ferreyra-Suarez et al., 2023). Rice bran protein (RBP) is a high-quality protein is characterised by a high lysine concentration, balanced amino acid composition and low allergenicity (Chalamaiah et al., 2017; Prakash & Ramaswamy, 1996); thus, RBP resources are increasingly valued. However, RBP combines through disulphide bonds, forms macromolecular complexes, and significantly reduces its ability to dissolve in water, even it has poor functional characteristics (Champagne et al., 2004); thus, RBP resource development is limited (Sophitha et al., 2021).
The preparation of protein fibrils could improve their gel formation, surface hydrophobicity, emulsification, interfacial activity, and functional properties. Thus, they have attracted the attention of the food industry for applications in food and materials (Jia et al., 2021). Protein fibrils are usually prepared from natural protein such as whey protein isolates, wheat flour proteins, and milk proteins by heating them at low pH (usually pH < 2.0), low ionic strength, and high temperature (>90 °C) (Rabe et al., 2020). During fibrils formation the internal protein bonds are broken, the structure of protein is unfolded, revealing the amino acid side chains that are part of the side chain amino/carboxylic group, as well as the hydrophobic side chains and hydroxyl groups (Liu et al., 2024). It is reported that protein fibrils have a high aspect ratio and provide better emulsification than rigidly bound spherical particles (Peng et al., 2017). It has been demonstrated that protein aggregates could form a more elastic interfacial film at the emulsion interface, thereby reducing surface tension. Protein fibrils are also commonly employed in the preparation of oil-in water (O/W) emulsions. Afkhami et al. investigated the preparation of O/W emulsions of soybean isolate protein fibrils with varying β-sheet structural contents and differing fibrils lengths. The sample with the highest β-sheet concentration and the longest fibrils length had the fastest adsorption kinetics at the oil/water interface, resulting in a decrease in droplet size, a more even distribution, and increased viscosity (Afkhami et al., 2023). Dong et al. examined the impact of fibrils alterations on the composition, capacity to form emulsions, and durability of emulsions made from egg white protein under various conditions. These findings indicate that the formation of fibrous structures was influenced by a range of factors. This includes measurements of pH, protein concentration, and treatment duration. At a pH of 2.0 and a protein concentration of 2% (wt/vol), the quantity of protein fibrils increased with increasing heating duration. The emulsified oil droplets with egg white protein fibrils are reduced in size, increase in surface potential and increase in stability (Dong et al., 2022).
An emulsion is a mixture of two or more liquids that do not mix together, where one liquid is dispersed in the other in the form of small droplets. These droplets are kept stable by a substance called an emulsifier. Double emulsions consist of three solutions and have smaller droplets interspersed within larger emulsion droplets (Sebben et al., 2021). Double emulsions can be classified into two distinct types: oil-in-water-in-oil and water-in-oil-in-water (W/O/W) emulsions. The preparation of double emulsions could be divided into two steps. The initial stage involves the preparation of the primary W1/O emulsion employing a lipophilic emulsifier. The second step involves the preparation of the W1/O/W2 emulsion using an alternative hydrophilic emulsifier (Wang et al., 2017). Its application in the food industry shows greater potential, as its unique physicochemical properties endow food with specific taste, texture, and nutritional functions. Furthermore, with the increasing consumer demand for health and the growing focus on sustainability, new formulations and innovative products based on emulsions continue to emerge. Owing to their distinctive structural characteristics, double emulsions are employed in a multitude of industrial applications, including the production of low-fat products, the improvement of sensory characteristics, the delivery of protective active ingredients, and the manufacture of cosmetics (Ding et al., 2019; Niyom et al., 2023). Researchers have utilised ethanol to encapsulate capsaicin within the interior aqueous phase of a double emulsion with an oil-in-water (W/O/W) structure. These findings indicated that encapsulation significantly increased the bioaccessibility of capsaicin and reduced gastrointestinal tissue irritation in mice (He et al., 2023). Compared with traditional single-layer emulsions, double emulsions are more complex and costly to prepare. They are also susceptible to decomposition under ambient pressure, which not only prevents internal droplets from aggregating but also prevents internal droplets from merging with the external solution (Muschiolik & Dickinson, 2017). There is considerable demand for emulsifiers that maintain emulsion stability.
This study aimed to analyse the influence of fibrils formation on the structure and W/O/W double emulsification characteristics of RBP via morphological, secondary structural, and molecular weight analyses. Also it examined the impact of the RBP fibrils structure on the stability of W/O/W double emulsions.
Materials and methods
Materials
Rice bran was acquired from Minyi Microbial Technology Development Company (Jilin, China). The protein content of the rice bran was quantified at 13.759% via the Kjeldahl method. The soybean oil was obtained from Yihai Jiali Jinlongyu Food Group Co., Ltd., located in Liaoning, China. T was obtained from Sigma–Aldrich (St. Louis, MO). The experiment utilised sodium dodecyl sulphate (SDS) (Coolaber Science & Technology). All the other compounds used were of analytical quality and sourced from China.
RBP extraction
Fresh rice bran was pulverised to a fine powder in a grinder and subsequently passed through an 80-mesh sieve. The rice bran powder was defatted by soaking it in hexane three times. The material was subsequently dried in an oven at 40 °C and stored in a refrigerator at −28 °C. For additional details on the extraction of RBP, please refer to the work of Tanger et al. (2020). The process involved dissolving skimmed rice bran in distilled water at a solid–liquid ratio of 1:10 (wt/vol). The pH was adjusted to 9.5 using a 2.0 M sodium hydroxide solution. The mixture was subsequently vigorously agitated in a water bath at 40 °C for 4 hr and then subjected to centrifugation at 4,000×g for 15 min. The liquid portion was collected, and the pH was adjusted to 4.5 using a solution of 1 M hydrochloric acid. The solution was subsequently forcefully stirred and then centrifuged at a force equivalent to 4,000 times the acceleration due to gravity for a period of 15 min, leading to the development of the protein precipitate. The precipitate subsequently underwent freeze drying.
RBP fibril preparation
Protein fibrils were created via the technique developed by Misumi et al. with some adjustments (Misumi et al., 2022). Dried RBP powder was subjected to milling and sieving. Dissolve 20 g of RBP in 500 ml of distilled water, allow it to fully hydrate, and prepare a protein solution with a concentration of 40 mg/ml. The RBP solution was adjusted to pH 2.0 with 5 M HCI, then loaded into sealed test tubes and placed in a 90 °C water bath for heating (0, 1, 2, 3, 4, 5, and 6 hr), for each corresponding heating time period, take out 50 ml of the protein solution while continuing uninterrupted heating, with a change in the temperature of the bath of less than 1 °C. The samples were subsequently subjected to thermal treatment in a water bath maintained at 90 °C for 0.5 hr. Next, the samples were subjected to cooling in an ice bath. Afterward, the samples were extracted from the heating equipment and cooled to ambient temperature via an ice–water bath. A portion of each sample was freeze-dried to remove water, while the remainder was subjected to the subsequent step of the experiment.
Morphology of fibrils revealed with transmission electron micrscopy
The protein fibrils structure was imaged via transmission electron micrscopy (TEM). The RBP fibrils solution was diluted 50-fold with phosphate buffer (pH 7.4，0.01 M). Subsequently, a carbon-coated copper grid was subjected to 10 μl of the dilution. After 2 min, the surplus solution was removed with filter paper. After air drying in a desiccator at 25 °C, the solution was treated with a 2% phosphotungstic acid stain, and images were captured with an accelerating voltage of 80 kV.
ThT fluorescence intensity analysis
Fluorescence measurements were employed as indicators of the extent of the cross-β-sheet structure, in accordance with the methodology of Khan et al. A solution of ThT at a concentration of 0.02 mg/ml was produced, 30 μl of PBP fibrils was added, and the reaction was monitored after 120 s (stored in the dark during the process). The fluorescence intensity of ThT was quantified via an F97 fluorescence spectrophotometer (Shanghai, China). The excitation wavelength was 460 nm, and the emission wavelength ranged from 440 to 520 nm. The width of the emission slit was set to 10 nm, and the magnitude of the voltage was adjusted to 800 mV.
Fourier transform infrared spectroscopy
The RBP fibrils powder, which was devoid of moisture, was combined with potassium bromide in a proportion of 1 part rice bran protein fiber structure aggregates (RBPA) powder to 100 parts potassium bromide. The resulting mixture was further pulverised to create translucent, slender slices. The test range spanned from 400 to 4,000 cm−1, with a resolution of 4 cm−1, and a total of 128 scans were conducted. The data were baseline corrected and smoothed via OMNIC software, and then the calculated secondary structure content was fitted to the amide I region via PeakFit 4.12.
Sodium dodecyl sulphate–polyacrylamide gel electrophoresis
The method of König et al. was used with minor modifications (König, 2019), with the lyophilised RBP fibrils powder (12 mg) dissolved in 1 ml of phosphate buffer solution (BPS) mixture and vortexed again for 1 min at 4 °C overnight. This method was modified slightly. The lyophilised powder of RBP fibrils (12 mg) was dissolved in 1 ml of PBS, vortexed for 1 min, hydrated overnight at 4 °C, and vortexed again for 1 min the following day. Fifty microliters of the supernatant was combined with 50 μl of upsampling buffer, which consisted of 0.1 mol/L Tris–HCl buffer (pH 6.8), 4% SDS, 2% 2-ME, 20% glycerol, and 0.4% bromophenol blue. Additionally, 10 μl of ammonium persulphate (APS) (10%) was completely mixed. The solution was heated in a water bath at boiling temperature for 5 min. Subsequently, 10 μl of the resulting mixture was introduced into the sample. The electrophoresis gels used were 5% concentrated gels and 12% separation gels. The electrophoresis system was installed, and electrode buffer was added. The voltage was initially adjusted to 80 V and maintained for 30 min. The voltage was subsequently increased to 120 V, and electrophoresis was stopped when the blue band was 1 cm from the bottom. First, the samples were fixed for 40 min with a fixing solution and subsequently dyed for 40 min with a dyeing solution. The samples were subsequently subjected to a decolourisation process, during which they were immersed in a decolourising solution for 5 hr. This was followed by replacement of the decolourising solution every hour. Next, the samples were stained with decolourising solution for 40 min.
Emulsifying activity index and emulsifying stability index
We adopted and slightly modified the approach of Agyare et al. (2009). A 15-ml sample of PBP fibrils was combined with 5 ml of soybean oil. The homogenisation process was conducted at 20,000 rpm for 1 min, after which the sample was immediately transferred to a graduated tube. At various time intervals, 50 μl of the Pickering emulsion was extracted from the lower layer, and an equal volume of the aqueous phase from the lower layer was collected in the same manner. These samples were then combined with 5 ml of 0.1% SDS. SDS was added to 100 ml of 0.01 mol/L phosphate buffer. The SDS and rice bran protein (RBP) fibrils were combined and agitated via a vortex shaker. The absorbance values were then measured at a wavelength of 500 nm and adjusted by subtracting the absorbance of a 0.1% SDS solution used as a blank. The emulsion activity index (EAI) and emulsion stability index (ESI) were computed via the following formulas:
The following variables are considered:
C0: Concentration of the RBP fibrils sample (g/ml).
A10: Absorbance value at an emulsion resting time of 10 min.
A0: Measurement of light absorption by the emulsion when it is at rest for 0 min.
ΔT: Emulsion standing time, 10 min.
Preparation of W/O/W emulsions with RBP fibrils
W/O/W emulsions were made via the techniques described by Paximada et al. (2021). The protein concentration of the RBP fibrils solution was 1% (wt/vol). Furthermore, the solution was agitated at ambient temperature for 2 hr. The solution was completely saturated with water in the internal aqueous phase, with a W/O/W volume ratio of 4:8:8. The internal aqueous phase was mixed with the oil phase and then blended at room temperature for 60 s in a high-speed homogeniser at 13,500 rpm to produce colostrum. The external aqueous phase was subsequently combined with the colostrum and subjected to homogenisation for 40 min at room temperature via a high-speed homogeniser operating at 9,000 rpm, resulting in the production of colostrum.
Emulsion properties and structural characterisation
Analysis of emulsion particle size and zeta potential
The particle size of an emulsion refers to the average diameter of the dispersed phase (typically droplets) within the emulsion and is one of the key parameters for evaluating the physical properties of the emulsion (Liu et al., 2021). The emulsions were analysed via an S3500 laser particle sizer (MICROTRAC, US). The solution was diluted with deionised water to achieve a concentration of 0.1% (wt/vol). The refractive index of the oil phase was measured as 1.465, whereas the refractive index of the aqueous phase was measured as 1.333. The average of three measurements was taken. The emulsions were subsequently analysed by a Zeta Sizer Nano Z90 particle size potentiostat following dilution by a factor of 125 with deionised water. The refractive coefficient was determined to be 1.450. The sample was allowed to reach thermal equilibrium for 2 min at 25 °C. The experiments were conducted in triplicate.
Percentage of adsorbed proteins
This study aimed to investigate the influence of RBP and RBP fibrils, which are used as emulsifiers, on double emulsions inside emulsions. This study examined how protein aggregation affects the composition of emulsion interfaces by quantifying the amount of protein adsorbed onto the surface (AP). The AP measurements were conducted in accordance with the methodology proposed by Shao et al. Shao & Tang (2014). The sample was prepared by introducing 20 ml of the solution. The emulsion was subjected to centrifugation at a rate of 4,000 revolutions per minute for a period of 10 min. The lower layer of the liquid, which was transparent and formed distinct layers, was subsequently meticulously removed via aspiration. The protein concentration was subsequently assessed via Koammas Brilliant Blue, and the AP (%) of the emulsion was computed.
where Cs, total protein content in the emulsion (mg/ml).
Cf, protein content of the aqueous phase after centrifugation (mg/ml).
Rheological properties
The rheological properties of the double emulsion were immediately examined after preparing the emulsion with a protein concentration of 24 mg/ml, diluted by the oil phase, and the dynamic and steady shear rheological properties of the emulsions were obtained via a hybrid rheometer (DHR-1 10nN.m-50mN.m; TA Instruments, New Castle, DE, USA), the test method of which is based on Janssen et al. (2020). A 40-mm diameter plate-plate geometry with a measuring gap of 1,000 μm. First, flow scan mode was used with a shear rate of 0.01–100 s-1, maintaining a temperature of 25 °C, and the viscosity of the emulsion was evaluated by calculating the ratio of 0.1–100 s.
For the dynamic oscillatory scanning test, a 40-mm diameter plate was used for the determination, and the gap was set at 1,000 μm. The emulsion to be tested was placed on the platform, and the plate was pressed down to the trim distance. The excess sample around the plate was scraped off, and silicone oil was used for sealing. At a strain value of 0.2% and a temperature of 25 °C, an angular frequency stigma was used to determine the energy storage modulus (G’) and the loss modulus (G”) of the emulsion.
Storage stability evaluation
An equal amount of emulsion was taken in a 10-ml centrifuge tube and placed upside down in front of a black background, and images of the apparent state of the emulsion were acquired at 0, 12, and 24 hr.
Microscopic imaging observation of the emulsion
A light microscope with a ×10 objective was used for observation. Prior to sampling, the solution was gently inverted three to five times. One droplet was mixed with 4 ml of water, resulting in a total volume of 1 ml. The emulsion was added to microscope slides and then overlaid with a glass coverslip. The studies were replicated five times, and only sample photographs are shown. The emulsion was sustained for 10 or 20 min for imaging.
Microstructure
In accordance with the methodology proposed by Yan et al. (2021), the emulsions were stained with minor adjustments. An SP-2 confocal laser scanning microscope (CLSM) was used to investigate the microstructure of the emulsions (Leica, Germany). The soybean oil was stained with Nile red at a concentration of 1 mg/ml in dimethyl sulphoxide, while the proteins were stained with Nile blue at a concentration of 1 mg/ml in Milli-Q water. The emulsion samples, which were labelled with fluorescent markers, were positioned on concave confocal microscope slides. They were then secured with a glass cover and captured via a 40× objective lens with oil immersion and a numerical aperture of 1.4. The pinhole diameter was kept constant at 1 Airy spot unit to effectively eliminate most of the light scattering. The excitation wavelengths employed for Nile red and Fast Green were 488 and 633 nm, respectively. The emission filter length for Nile red was adjusted to 555–620 nm, whereas that for Nile blue was configured to 660–710 nm.
Statistical analysis
The studies were conducted three times, and one-way analysis of variance was performed via SPSS 26. The Duncan model was used to examine potential disparities, with a significance level of p < .05. Line, bar, and infrared graphs were generated via Origin 9.1 software. Image stitching was performed via PowerPoint 2016 and Photoshop software.
Results and discussion
Morphology of fibrils formation of RBP
TEM was employed to elucidate the RBP fibrils structure formation process following heat treatment (0, 2, 4, and 6 hr). The unheated RBP exhibited a large agglomerate molecular structure (Figure 1A). The formation of the structure was driven mainly by hydrophobic interactions and hydrogen bonding. The number of large agglomerates was significantly lower after the RBP was heated for 2 hr (Figure 1B), indicating that the enhancement of the hydration effect was significant, which contributed to the dissociation of agglomerate groups by heating and the exposure of the hydrophobic groups, thereby increasing the number of protein–protein interactions. After the RBP was heated for 4 hr, the number of small aggregated fragments, as well as fibrils of shorter length (Figure 1C), was related to disulphide bonds, hydrogen bonds, and other interactions during aggregation (Zaman et al., 2014). After 6 hr heating, it became evident that the RBP had undergone aggregation, with the majority of the resulting fibril structures exhibiting an elongated and straight configuration (Figure 1D). The RBP was heated at pH 2.0, where upon protein aggregation was driven by electrostatic interactions. This process is consistent with the “nucleation growth theory” of fibrils structures (Kashchiev, 2015). The structure of the fibrils formed was also in accordance with the findings of Kutzli et al. The fibrils presented a high aspect ratio, with the formation of these fibrils being attributed to the use of cannabis seed proteins (Kutzli et al., 2022).
Transmission electron micrscopy images of rice bran protein during the fibrillar trembling. (A) Heating for 0 hr, (B) heating for 2 hr, (C) heating for 4 hr, (D) heating for 6 hr.
Thioflavin T fluorescence analysis
The ability of protein fibrils to selectively attach to the β-sheet conformation makes them a highly efficient and responsive technique for tracking the development of amyloid fibrils (Sulatsky et al., 2020). Figure 2 displayed the relationship between the fluorescence intensity of the RBP and the duration of heating, which ranged from 0 to 6 hr. The fluorescence intensity progressively increased as the fibrils creation process advanced due to the development of the β-sheet structure. This suggested that heating derived the formation and growth of the fibrils structure. The low fluorescence intensity of the unheated samples could be attributed to the fact that thioflavin T (ThT) does not bind to amorphous aggregates. Following 2 hr heating period, compared to the unheated RBP there was a small increase in the brightness of the fluorescence, which was in accordance with the TEM findings. The increase in ThT binding could be ascribed to the disruption of the aggregate framework caused by heating at pH 2.0, leading to the creation of smaller oligomers that were capable of binding with Th T (Qin et al., 2017). The inset in Figure 2 showed that the trend of increasing fluorescence intensity with time followed by a rapid increase was consistent with the trend of lagged changes in nucleated self-assembly reactions (Zhang & Schmit, 2017). Lei & Ma (2021) reported the same findings. The observed alterations in the fibrils structure of fibrils samples derived from wheat gluten proteins align with the results of the current investigation.
Thioflavin T (ThT) spectroscopic profiles of rice bran protein during the fibrillar trembling. The inset indicates the maximum fluorescence intensity of ThT as a function of heating time.
Secondary structure of RBP during the fibrils formation
The amide I band, ranging from 1,600 to 1,700 cm−1, is the primary distinctive protein band observed via Fourier theorem infrared spectroscopy. During the synthesis of protein fibrilss, the secondary structures were determined to be α-helices (1654 cm−1), β-sheets (1,618, 1,629–1,637, and 1,679–1,697 cm−1), random coils (1647 cm−1), and β-turns (1,664–1,672 cm−1). The alterations in protein secondary structure during the process of protein fibrils synthesis were displayed in Table 1. The content of the α-helix structure gradually decreased, indicating the creation of an organised β-sheet structure and rearrangement of hydrogen bonds during the process of heating. The hydrogen bonds in the α-helix can stably reinforce the local structure of polypeptides without interference from side chains, providing proteins with rigidity and elasticity. The α-helix tends to arrange hydrophobic and hydrophilic side chains in a manner that helps proteins achieve the correct conformation in aqueous or membrane environments. In contrast, the β-sheet structure, stabilised by parallel or antiparallel hydrogen bonds, further enhances the overall mechanical strength and tensile properties of proteins, as exemplified in silk fibroin (Promsuk et al., 2024). In amyloid fibers and many structural proteins, β-sheets are key components in forming crystalline cores. These findings aligned with prior research conducted by Xia et al. on alterations in the secondary structure that transpire during the fibrillation of yeast proteins (Xia et al., 2022).
The secondary structure of rice bran protein during the fibrillar trembling.
| Heat time (h) | β-Sheet (%) | Random coil (%) | α-Helix (%) | β-Turn (%) |
|---|---|---|---|---|
| 0 | 36.81 ± 0.52g | 19.85 ± 0.08a | 31.16 ± 31.16a | 12.18 ± 0.27c |
| 1 | 38.27 ± 0.17f | 19.83 ± 0.15a | 29.66 ± 29.66b | 12.24 ± 0.19c |
| 2 | 38.99 ± 0.16e | 19.79 ± 0.12a | 28.64 ± 28.64c | 12.58 ± 0.18bc |
| 3 | 39.56 ± 0.25d | 19.91 ± 0.06a | 27.62 ± 27.62d | 12.91 ± 0.09b |
| 4 | 40.12 ± 0.06c | 19.88 ± 0.10a | 27.24 ± 27.24d | 12.76 ± 0.14b |
| 5 | 40.6 ± 0.05b | 19.84 ± 0.10a | 26.83 ± 26.83e | 12.73 ± 0.3b |
| 6 | 41.96 ± 0.09a | 19.93 ± 0.14a | 24.72 ± 24.72f | 13.39 ± 0.41a |
| Heat time (h) | β-Sheet (%) | Random coil (%) | α-Helix (%) | β-Turn (%) |
|---|---|---|---|---|
| 0 | 36.81 ± 0.52g | 19.85 ± 0.08a | 31.16 ± 31.16a | 12.18 ± 0.27c |
| 1 | 38.27 ± 0.17f | 19.83 ± 0.15a | 29.66 ± 29.66b | 12.24 ± 0.19c |
| 2 | 38.99 ± 0.16e | 19.79 ± 0.12a | 28.64 ± 28.64c | 12.58 ± 0.18bc |
| 3 | 39.56 ± 0.25d | 19.91 ± 0.06a | 27.62 ± 27.62d | 12.91 ± 0.09b |
| 4 | 40.12 ± 0.06c | 19.88 ± 0.10a | 27.24 ± 27.24d | 12.76 ± 0.14b |
| 5 | 40.6 ± 0.05b | 19.84 ± 0.10a | 26.83 ± 26.83e | 12.73 ± 0.3b |
| 6 | 41.96 ± 0.09a | 19.93 ± 0.14a | 24.72 ± 24.72f | 13.39 ± 0.41a |
Notes: a,b,c:Indicating significant differences in data (p < 0.05).
The secondary structure of rice bran protein during the fibrillar trembling.
| Heat time (h) | β-Sheet (%) | Random coil (%) | α-Helix (%) | β-Turn (%) |
|---|---|---|---|---|
| 0 | 36.81 ± 0.52g | 19.85 ± 0.08a | 31.16 ± 31.16a | 12.18 ± 0.27c |
| 1 | 38.27 ± 0.17f | 19.83 ± 0.15a | 29.66 ± 29.66b | 12.24 ± 0.19c |
| 2 | 38.99 ± 0.16e | 19.79 ± 0.12a | 28.64 ± 28.64c | 12.58 ± 0.18bc |
| 3 | 39.56 ± 0.25d | 19.91 ± 0.06a | 27.62 ± 27.62d | 12.91 ± 0.09b |
| 4 | 40.12 ± 0.06c | 19.88 ± 0.10a | 27.24 ± 27.24d | 12.76 ± 0.14b |
| 5 | 40.6 ± 0.05b | 19.84 ± 0.10a | 26.83 ± 26.83e | 12.73 ± 0.3b |
| 6 | 41.96 ± 0.09a | 19.93 ± 0.14a | 24.72 ± 24.72f | 13.39 ± 0.41a |
| Heat time (h) | β-Sheet (%) | Random coil (%) | α-Helix (%) | β-Turn (%) |
|---|---|---|---|---|
| 0 | 36.81 ± 0.52g | 19.85 ± 0.08a | 31.16 ± 31.16a | 12.18 ± 0.27c |
| 1 | 38.27 ± 0.17f | 19.83 ± 0.15a | 29.66 ± 29.66b | 12.24 ± 0.19c |
| 2 | 38.99 ± 0.16e | 19.79 ± 0.12a | 28.64 ± 28.64c | 12.58 ± 0.18bc |
| 3 | 39.56 ± 0.25d | 19.91 ± 0.06a | 27.62 ± 27.62d | 12.91 ± 0.09b |
| 4 | 40.12 ± 0.06c | 19.88 ± 0.10a | 27.24 ± 27.24d | 12.76 ± 0.14b |
| 5 | 40.6 ± 0.05b | 19.84 ± 0.10a | 26.83 ± 26.83e | 12.73 ± 0.3b |
| 6 | 41.96 ± 0.09a | 19.93 ± 0.14a | 24.72 ± 24.72f | 13.39 ± 0.41a |
Notes: a,b,c:Indicating significant differences in data (p < 0.05).
Composition of RBP during the fibrils formation
The molecular weights of the RBPs ranged primarily from 10 to 75 kDa, as shown in Figure 3. The bulk of these weights corresponded to globulin subunits, albumin subunits, glutenin acidic subunits, and glutenin basic subunits and proprotein subunits. After heating for 6 hr, the RBP with a molecular weight of 15 kDa (acidic) subunits disappeared, indicating that the RBP was hydrolysed into smaller structures. Consequently, at pH 2.0, the acidic subunit was temperature sensitive and readily decomposes to form smaller polypeptides, a process that was conducive to the fibrils formation process. Additionally, Tang et al. reported the appearance of small molecular weight fragments following the extreme heating of kidney bean 7S globulin (Mw < 10 kDa at > 360 min) (Tang et al., 2010).
Sodium dodecyl sulphate polyacrylamide gel electrophoresis of rice bran protein during the fibrillar trembling. M = Maker; A–G = heating for 0–6 hr).
EAI and ESI of RBP during the fibrils formation
Figure 4A displayed the EAI and ESI of the RBP fibrils. The EAI quantifies the ability of protein to adhere to the interface between water and oil, as well as its ability to resist flocculation and aggregation caused by gravity and electrostatic interactions. As illustrated in Figure 4A, the PBP fibrils prepared following a heating period of 6 hr presented remarkable values for both the EAI and ESI. The EAI increased to 234.66 m2/g, representing a 33.09% increase compared with that of the natural RBP. Additionally, the ESI was increased by 115.44% in comparison with that of the natural RBP. The observed increase in EAI can be attributed to the disturbance of the initial protein structure caused by heat, leading to the exposure of additional hydrophobic groups. These groups have the ability to move quickly at the boundary between oil and water, which helps in the creation of protein films on the surface during the process of emulsion. The observed increase in the ESI might be attributed to the formation of a thicker and more resilient protein interfacial membrane at the water–oil interface, which reduced the interfacial tension. Notably, Carter et al. also demonstrated that fibrillated lentil proteins can be employed to stabilise emulsions over an extended period (Wynnychuk et al., 2021).
Changes in the effects of RBPAs on lotion properties (A) Changes in the emulsion activity index and emulsion stability index of RBPAs with heating time; (B) Changes in the particle size of the Wp/O/Wp lotion prepared with RBPAs; (C) Changes in the potential of the Wp/O/Wp lotion prepared with RBPAs; (D) Percentages of adsorbed proteins.
Analysis of emulsion particle size and zeta potential
The size of the emulsion droplets had an impact on the overall performance of the emulsion. Figure 4B and C showed that the emulsion droplet sizes stabilised by the formation of protein aggregates at various stages of RBP fibrils preparation. Compared with that of natural RBP emulsions, the droplet size of protein emulsions in which fibrils formation occurred was significantly reduced. The smallest emulsion particle size stabilised by RBP fibrils with an elongated fibrous structure formed by heating for 6 hr was 1171.6 nm, Compared to the emulsions stabilised by RBP with a heating time of 0 hr (1691.6 nm), the droplet size decreased by 69.26%, showing a significant change. It has been observed that emulsion droplets prepared from fibrils-structured proteins are smaller in size (Lee et al., 2023). Thus, RBP fibrils might form a thick layer with elasticity at the emulsion interface, thereby reducing the droplet size. Moreover, as the β-sheet structure and fibrils length increased, the emulsion droplet size decreased.
The ζ-potential could be employed to predict the stability of an emulsion system. Emulsions with higher ζ-potentials are more stable because of the presence of an electric charge within the emulsion, which generates high repulsive forces between droplets, preventing droplet aggregation. For the emulsion system, stability was achieved when the electric potential was above 10 mV or below −10 mV. As shown in Figure 4C, the emulsion ζ-potential was consistently below −10 mV when the RBP was subjected to heating 2 hr later. With prolonged heating, the ζ-potential initially decreased but subsequently increased. The lowest ζ-potential was −17.5 mV for the heating period of 4 hr. The pH of RBPA solution was 2.0, which was below the isoelectric point, the majority of the protein charge originated from the amino groups. Consequently, the majority of the charge was derived from the amino group of the protein (Mohammadian et al., 2018). Fibrils formation resulted in the exposure of the amino groups encapsulated within the RBP without inducing alterations in hydrophobicity or ζ-potential.
AP of RBP during the fibrils formation
Figure 4D showed the stable W/O/W of the RBP for different heating hours. The hydrophobicity of the protein and its flexibility at the oil–water interface directly affect the adsorption behaviour of the protein. The AP of the natural RBP was 30.0647%, and it was increased to 47.2834% after 1 hr of heating. Fibrils formation processing revealed the hydrophobic amino group within the protein molecule and enhanced its flexibility by revealing its molecular structure. This change was consistent with the abovementioned causes of a decrease in the ζ potential. The hydrophobic groups that were visible interacted with the oil droplets, facilitating the adsorption of the protein at the interface between the oil and water. After heated for 6 hr, the AP reached its peak value of 62.3547%. These findings corroborated the results of Wei et al. (2019).
Rheological analysis of RBP during the fibrils formation
Figure 5 showed the stable emulsion fluidity and dynamic viscoelasticity of the aggregates obtained from RBP at different heating times. As shown in Figure 5A, all emulsions exhibited shear-thinning flow behaviour, suggesting that the interaction between protein molecules adsorbed at the oil–water interface in the emulsion maintained emulsion stability and prevented droplet flocculation. At the initial shear rate of the emulsion, except for the protein-stable emulsion obtained by heating for 5 or 6 hr, there was no obvious difference in the other viscosities, and the emulsion at this time had less aggregation, so the particle size of the droplets in the emulsion was small, and the spatial repulsion between the droplets was high. With increasing shear rate, the viscosity of the emulsion decreased significantly, indicating that the shear damaged the protein interface membrane between the oil and water in the emulsion. However, after 5–6 hr of acid heat treatment of RBP, the decrease in the viscosity of the stabilised emulsions was significantly reduced. Moreover, with the increase in shear rate, the viscosity of the W/O/W emulsions was higher compared to those prepared with RBP samples subjected to shorter heating times. At lower frequency scans, the G” of the double emulsion was greater than the G’, indicating that the emulsion was more viscous at this time. With increasing frequency, both G’ and G” increased, and G’ was greater than G”, indicating that the unwanted network structure began to appear in the small emulsion at higher frequencies and that the viscous properties of the fluid gradually changed to the elastic structure of the solid state. As shown in Figure 5H, the emulsion samples subjected to protein heat treatment for 6 hr were the first to change to a weak gel state, which could be explained by the amyloid fibrillar structural protein formed by RBP after acid heat treatment, and the gel properties improved. This phenomenon has also been confirmed by Wang et al. (2024). Another reason might be that some of the microdroplets that did not form a double emulsion during the shearing process wrap large droplets, achieving the Pickering effect, strengthening the structure of the emulsion, and promoting the gel structure of the emulsion. This phenomenon, although less common, has been proven feasible by Apostolidis et al. (2024).
Effect of RBPA on the rheological properties of emulsions during the fibrillar trembling. (0, 1, 2, 3, 4, 5, and 6 hr) (A) Apparent viscosity (η) versus shear rate (γ), (B–H) Viscoelastic properties of RBPA-stabilised lotion obtained with different treatment times.
Storage stability of emulsion
Storage stability evaluation
Figure 6 showed that the newly prepared W/O/W RBP emulsions prepared at different heating times were stable and maintained the base phase of the double emulsion. After 12 hr, all of the emulsions delaminated, except for the samples heated for 5 and 6 hr, which did not show any obvious delamination. The main reason for this delamination was that RBP, the coemulsifier of the double emulsion, lacked proper equilibrium between its affinity for lipids and its affinity for water. The emulsifying ability of RBP is improved by fibrils formation, so the amount of water phase precipitated at the bottom is almost zero after 24 hr, although insignificant delamination can also be observed.
Appearance characterisation of w/o/w lotion prepared by rice bran protein with different structures (A) heat treatment for 0 hr, (B) Heat treatment for 12 hr, (C) heat treatment for 24 hr.
Microscopic imaging observation of the emulsion
As illustrated in Figure 7, the W/O/W emulsions prepared from RBP produced at different heating times demonstrated that the emulsion heated for 6 hr exhibited the best performance following stabilisation for 20 min. This was evidenced by the observation that the interfacial film thickness between water and oil was the greatest. This represented a clear advantage over emulsions stabilised with natural RBP. The improved emulsification capability could be ascribed to the reinforcement of hydrophobic interactions among protein molecules, disulphide bonding, and hydrogen bonding, leading to the creation of a more enduring water–oil–protein interfacial film (Mishchuk et al., 2004). These results align with the conclusions of prior research on the effectiveness and efficiency of RBP fibrils.
Microscopic image of water-in-oil-in-water bilayer emulsion prepared by rice bran protein with different acid heat treatment time (A–G) heating 0–6 hr; (A–G) heating for 1: image under microscope of newly prepared emulsion; 2: image under microscope after 10 min of emulsion; 3: image under microscope after 20 min of emulsion.
CLSM of RBP during the fibrils formation
CLSM was used to examine the microstructure of the W/O/W emulsions stabilised by RBP and RBP fibrils during the fibrils formation. This observation was shown in Figure 8. The oil phase is indicated in green, while RBP and RBP fibrils are indicated in red. As illustrated in the accompanying figure, the RBP with slender fibrils, which was produced by heating for 6 hr, formed a dense interfacial layer around the droplets. The viscosity test demonstrated that RBP fibrils increased the viscosity of the emulsion. A dense and thick protein interfacial coating might serve as a spatial barrier, thereby resisting the instability of phase separation and aggregation (Atarian et al., 2019). In line with the results of our prior investigation, Fu et al. similarly reported that the creation of stable emulsions using protein fibrils architectures is a highly effective approach (Fu et al., 2023).
Images of confocal laser scanning microscope of RBPAs during the fibrillar trembling. (A) Unheated rice bran protein. (B) Lotion prepared after heating for 6 hr.
Conclusion
To summarise, the most favourable conditions for the creation of RBP fibrils were identified as heating the RBP at a concentration of 1% at a pH of 2.0 and a temperature of 90 °C for a duration of 6 hr. The formation of RBP fibrils was found to follow the nucleation–growth theory and was accompanied by protein denaturation and hydrolysis of the acidic subunits, as well as the generation of new aggregated structures. The exposure of the RBP hydrophobic group during heat treatment influenced the adsorption behaviour of the protein at the oil–water interface. Compared with RBP heated for 0 hr, the particle size of RBP fibrils emulsion in the emulsion decreased by 69.26%, shear viscosity increased significantly, the thickness of RBP fibril at the oil–water interface increased, and the W/O/W stability was significantly improved (p < .05). The increase in emulsion characteristics was directly linked to the interfacial features of the RBP fibrils. The impact of protein fiber structure on the properties of emulsions is a multifaceted topic, involving the molecular structure of the protein itself, its aggregation behaviour, and its interactions with other components in the emulsion. This influence is typically reflected in the emulsion’s stability, viscosity, interfacial properties, and long-term physical and chemical performance. In addition, proteins in the continuous phase display a low level of gelation. This further supports the development and utilisation of emulsion resources in industries such as food and cosmetics. It provides ideas for the further development and utilisation of plant processing by-products. Emulsion stabilisers derived from plant proteins might be employed by the food and cosmetic industries as sustainable, environmentally friendly, and cost-effective alternatives to traditional stabilisers.
Author contributions
Keyang Sun (Formal analysis [equal], Investigation [equal], Methodology [equal], Validation [equal], Writing—original draft [equal], Writing—review & editing [equal]), Yuzhe Gao (Conceptualization [equal], Funding acquisition [equal], Project administration [equal], Supervision [equal]), Yuan Yuan (Formal analysis [equal], Investigation [equal], Methodology [equal], Validation [equal], Writing—original draft [equal]), Yongping Xu (Formal analysis [equal], Investigation [equal], Methodology [equal], Validation [equal], Writing—original draft [equal]), Xiaojie Gong (Formal analysis [equal], Investigation [equal], Writing—original draft [equal]), Wanyue Jiang (Formal analysis [equal], Writing—original draft [equal], Writing—review & editing [equal]), Junqi Pang (Formal analysis [equal]), Shuying Li (Formal analysis [equal]), Yuzhe Gao (Conceptualization [equal], Data curation [equal], Funding acquisition [equal], Project administration [equal], Resources [equal], Supervision [equal]), Qingyu Yang (Formal analysis [equal], Funding acquisition [equal], Project administration [equal], Resources [equal], Supervision [equal])
Funding
This work was supported by the Natural Science Foundation of Liaoning Province (No. 2022-MS-307) and Graduate Research Project of Basic Scientific Research Business Fee of Liaoning Provincial Universities (SYNU SJ2024016).
Conflicts of interest
The authors declare no conflicts of interest.
Acknowledgements
The author guarantees that the data provided in the manuscript is usable. The author would like to give their heartfelt thanks to all the people who helped them with this paper.
References
Author notes
K.S. and Y.Y. contributed equally.
