A protein amyloid fiber-pyrazine aroma compound composite material and a preparation method and application thereof

Abstract

The present application belongs to the technical field of aroma substance stabilization, and provides a protein amyloid fiber-pyrazine aroma substance composite material, a preparation method and application thereof. The composite material comprises protein amyloid fiber and pyrazine aroma substance; the protein amyloid fiber comprises one or more of soybean protein amyloid fiber, pea protein amyloid fiber, rice protein amyloid fiber and wheat protein amyloid fiber. The present application introduces protein amyloid fiber, and the protein amyloid fiber and pyrazine aroma substance are combined to construct the most abundant intermolecular interaction network through hydrogen bond, van der Waals force and hydrophobic interaction, not only the number of effective combination sites is significantly increased, but also specific Pi-Sigma interaction is formed; when the protein amyloid fiber-pyrazine aroma substance composite material is applied to plant-based meat products, the combination sites with the plant-based meat products are more, and the combination and stability of the pyrazine aroma substance are improved.

Description

Technical Field

[0001] This invention relates to the field of aroma substance stabilization technology, and in particular to a protein amyloid fiber-pyrazine aroma substance composite material, its preparation method and application. Background Technology

[0002] With increasing global focus on health, environment, and animal welfare, plant-based meat products are rapidly becoming ideal alternatives to traditional meat. However, the lack of realistic flavor simulation, especially the absence of a distinctive "meaty aroma," remains a key sensory bottleneck limiting consumer acceptance. Pyrazine compounds, particularly alkylpyrazines, are key products of the Maillard reaction, imparting typical roasted and nutty aromas to meat products and are considered core flavor compounds for mimicking meat's characteristics. However, pyrazine compounds are highly volatile and chemically unstable, easily lost or experiencing flavor imbalances during the high-temperature processing and long-term storage of plant-based meat products. Therefore, constructing efficient flavor stabilization delivery carriers to effectively retain and control the release of pyrazine aroma compounds is a common challenge that urgently needs to be addressed to improve the sensory quality of plant-based meat products. Summary of the Invention

[0003] In view of this, the purpose of this invention is to provide a protein amyloid fiber-pyrazine aroma compound, its preparation method, and its application. The introduction of protein amyloid fibers in this invention improves the stability of the pyrazine aroma compounds.

[0004] To achieve the above-mentioned objectives, the present invention provides the following technical solution: This invention provides a protein amyloid fiber-pyrazine aroma compound composite material, comprising protein amyloid fibers and pyrazine aroma compounds; The protein amyloid fibers include one or more of soybean protein amyloid fibers, pea protein amyloid fibers, rice protein amyloid fibers, and wheat protein amyloid fibers; The pyrazine aroma compounds include one or more of 2,5-dimethylpyrazine, 2,6-dimethylpyrazine, 2,3-dimethylpyrazine, 2-methylpyrazine, and 2,3,5-trimethylpyrazine.

[0005] Preferably, the ratio of the protein amyloid fiber to the pyrazine aroma substance is 0.5~1.5g:0.05~0.1mmol.

[0006] Preferably, the method for preparing the protein amyloid fibers includes the following steps: The protein powder and water are first mixed, and the resulting first mixture is adjusted to be acidic. Hydration and fibrosis are then carried out sequentially to obtain the protein amyloid fibers.

[0007] Preferably, the mass ratio of the protein powder to water is 1:9~11, and the first mixing time is 10~14h.

[0008] Preferably, the acidic pH value is 1.5~2.5, the hydration time is 20~35 min, the fibrosis temperature is 80~90℃, and the time is 6~20 h.

[0009] This invention also provides a method for preparing the protein amyloid fiber-pyrazine aroma compound described in the above technical solution, comprising the following steps: Amyloid fibers were dispersed in a buffer solution, and then pyrazine aroma compounds were added. The resulting mixture was then equilibrated to obtain the amyloid fiber-pyrazine aroma compound composite material.

[0010] Preferably, the buffer solution is a phosphate buffer solution with a concentration of 10 mmol / L and a pH value of 6.5 to 7.5.

[0011] Preferably, the mass concentration of amyloid fibers in the mixture is 0.05~0.15%, and the concentration of pyrazine aroma compounds is 0.05~0.10 mmol / L.

[0012] Preferably, the equilibrium temperature is 37°C, the time is 15~20h, and the equilibrium is carried out under light-protected conditions.

[0013] The present invention also provides the application of the protein amyloid fiber-pyrazine aroma compound material described in the above technical solution or the protein amyloid fiber-pyrazine aroma compound material prepared by the preparation method described in the above technical solution in plant-based meat products.

[0014] This invention provides a protein amyloid fiber-pyrazine aroma compound composite material.

[0015] This invention introduces amyloid fibers, which combine with pyrazine aroma compounds through hydrogen bonds, van der Waals forces, and hydrophobic interactions to form the richest intermolecular interaction network. This not only significantly increases the number of effective binding sites but also forms specific Pi-Sigma interactions. When the amyloid fiber-pyrazine aroma compound is applied to plant-based meat products, it has a high binding site with the plant-based meat products, which improves the binding force and stability of pyrazine aroma compounds. Attached Figure Description

[0016] Figure 1The diagram shows the structural features of SAFs, PAFs, RAFs, and GAFs in Example 1. In this diagram, A is a transmission electron microscope image with a scale bar of 200 nm; B is the thioflavone T fluorescence curve; C is the X-ray diffraction pattern; D is the Fourier transform infrared spectrum; E is the synchronous fluorescence spectrum with Δλ=15 nm; F is the synchronous fluorescence spectrum with Δλ=60 nm; and G is the ultraviolet second derivative spectrum. SAFs: soybean protein amyloid fibers; SPI: soybean protein isolate; PAFs: pea protein amyloid fibers; PPI: pea protein isolate; RAFs: rice protein amyloid fibers; RP: rice protein; GAFs: gluten protein amyloid fibers; and GP: gluten protein. Figure 2 The figures show the dimensions of different protein-derived amyloid fibers in Example 1. A is the length histogram of SAFs, B is the diameter histogram of SAFs, C is the aspect ratio histogram of SAFs; D is the length histogram of PAFs, E is the diameter histogram of PAFs, F is the aspect ratio histogram of PAFs; G is the length histogram of RAFs, H is the diameter histogram of RAFs, I is the aspect ratio histogram of RAFs; J is the length histogram of GAFs, K is the diameter histogram of GAFs, and L is the aspect ratio histogram of GAFs. Figure 3 The image shows the structural features of SAFs, PAFs, RAFs and GAFs in Example 1, where A represents the thiol content, B represents the surface hydrophobicity, and C represents the SDS-PAGE image. Figure 4 The physicochemical properties characterization diagrams of SAFs, PAFs, RAFs and GAFs in Example 1 are shown, where A is the Zeta potential, B is the particle size and turbidity, C is the polydispersity index (PDI), D is the solubility, E is the thermogravimetric curve, F is the storage modulus (G'), G is the loss modulus (G''), H is the apparent viscosity, and I is the scanning electron microscope image with a scale bar of 500 μm. Figure 5 The diagram shows the aroma binding capacity of SPI, SAFs, PPI, PAFs, RP, RAFs, GP and GAFs in Example 1, where A is the aroma binding rate, B is the aroma binding thermal stability and C is the thermal analysis diagram. Figure 6 The images show the characterization of soy protein isolate and soy protein amyloid fibers in Example 2. In the images, A is a TEM micrograph with a scale bar of 200 nm, B is a ThT fluorescence measurement, C is an XRD pattern, D is an FTIR spectrum, E is a synchronous fluorescence spectrum at Δλ=15 nm, F is a synchronous fluorescence spectrum at Δλ=60 nm, and G is a second derivative ultraviolet spectrum. Figure 7The physicochemical properties characterization diagrams of SPI and SAFs in Example 2 are shown, where A represents the total thiol and free thiol content, B represents the surface hydrophobicity, and C represents the SDS-PAGE spectrum. Figure 8 The physicochemical and structural characteristics of SPI and SAFs in Example 2 are shown below, where A is the Zeta potential, B is the particle size and turbidity, C is the solubility, D is the polydispersity index (PDI), E is the thermogravimetric analysis (TGA) curve, F is the storage modulus (G'), G is the loss modulus (G''), H is the apparent viscosity, I is the scanning electron microscope (SEM) image with a scale bar of 200 μm, and J is the fiber yield. Figure 9 The interaction characterization pathways between SAFs and 2-methylpyrazine, 2,5-dimethylpyrazine and 2,3,5-trimethylpyrazine in Example 2 are shown in Figure 2. In the figure, A represents the binding ability, B represents the thermal stability curve, C represents the thermogram visualization, and D represents the UV-Vis absorption spectrum. Figure 10 The images show the structural characterization of pea protein isolate (PPI) and pea protein isolate amyloid fibers (PAFs) in Example 3. In the images, AC are TEM images with a scale bar of 200 nm, D is ThT fluorescence detection, E is XRD pattern, F is FTIR spectrum, G is synchronous fluorescence spectrum with Δλ=15 nm, H is synchronous fluorescence spectrum with Δλ=60 nm, and I is UV second derivative spectrum. Figure 11 The diagrams show the dimensions of PAFs-6, PAFs-12, and PAFs-18 in Example 3. Specifically, A is the length distribution histogram of PAFs-6, B is the length distribution histogram of PAFs-12, and C is the length distribution histogram of PAFs-18; D is the diameter distribution histogram of PAFs-6, E is the diameter distribution histogram of PAFs-12, F is the diameter distribution histogram of PAFs-18, G is the aspect ratio distribution histogram of PAFs-6, H is the aspect ratio distribution histogram of PAFs-12, and I is the aspect ratio distribution histogram of PAFs-18. Figure 12 The images show the physicochemical properties of PPI and PAFs in Example 3, where A represents the total and free thiol content, B represents the surface hydrophobicity, and C represents the SDS-PAGE image. Figure 13 The physicochemical and structural characteristics of PPI and PAFs in Example 3 are shown below. A represents Zeta potential, B represents particle size and turbidity, C represents solubility, D represents polydispersity index, E represents TGA curve, F represents storage modulus (G'), G represents loss modulus (G''), H represents apparent viscosity, I represents SEM image with a scale bar of 500 μm, and J represents fiber yield. Figure 14The binding capacity of PAFs to 2,3-dimethylpyrazine, 2,6-dimethylpyrazine, and 2,5-dimethylpyrazine in Example 3 is characterized by A, B, C, D, and D respectively. Figure 15 The synchronous fluorescence spectra of PAFs bound to different concentrations of pyrazine isomers in Example 3 are shown below. A represents the synchronous fluorescence spectrum of PAFs bound to 2,5-dimethylpyrazine at Δλ=15nm; B represents the synchronous fluorescence spectrum of PAFs bound to 2,6-dimethylpyrazine at Δλ=15nm; C represents the synchronous fluorescence spectrum of PAFs bound to 2,3-dimethylpyrazine at Δλ=15nm; D represents the synchronous fluorescence spectrum of PAFs bound to 2,5-dimethylpyrazine at Δλ=60nm; E represents the synchronous fluorescence spectrum of PAFs bound to 2,6-dimethylpyrazine at Δλ=60nm; and F represents the synchronous fluorescence spectrum of PAFs bound to 2,3-dimethylpyrazine at Δλ=60nm. Figure 16 The three-dimensional fluorescence spectra of PAFs combined with pyrazine isomers with different methyl positions in Example 3 are shown below. A is the three-dimensional fluorescence spectrum without pyrazine aroma substances, B is the three-dimensional fluorescence spectrum with 2,5-dimethylpyrazine added, C is the three-dimensional fluorescence spectrum with 2,3-dimethylpyrazine added, and D is the three-dimensional fluorescence spectrum with 2,6-dimethylpyrazine added. Figure 17 The images show the intrinsic fluorescence spectra of PAFs after binding with three different concentrations of pyrazine isomers in Example 3. In the images, A is 298 K, B is 308 K, C is 318 K, D is a Stern-Volmer plot, and E is a double logarithmic plot. Figure 18 This is a molecular docking diagram of PAFs with pyrazine aroma compounds at different methyl positions in Example 3; Figure 19 This is a schematic diagram illustrating the potential binding mechanism between PAFs and pyrazine aroma compounds with different methyl positions in Example 3; Different lowercase letters "ad" indicate significant differences between groups (p<0.05). Detailed Implementation

[0017] This invention provides a protein amyloid fiber-pyrazine aroma compound composite material, comprising protein amyloid fibers and pyrazine aroma compounds; The protein amyloid fibers include one or more of soybean protein amyloid fibers, pea protein amyloid fibers, rice protein amyloid fibers, and wheat protein amyloid fibers; The pyrazine aroma compounds include one or more of 2,5-dimethylpyrazine, 2,6-dimethylpyrazine, 2,3-dimethylpyrazine, 2-methylpyrazine, and 2,3,5-trimethylpyrazine.

[0018] In this invention, the preferred ratio of the protein amyloid fiber to the pyrazine aroma substance is 0.5~1.5g:0.05~0.1mmol, more preferably 1g:0.05~0.1mmol, and specifically preferably 1g:0.05mmol, 1g:0.06mmol, 1g:0.07mmol, 1g:0.08mmol, 1g:0.09mmol or 1g:0.1mmol.

[0019] In this invention, the method for preparing the protein amyloid fibers preferably includes the following steps: The protein powder and water are first mixed, and the resulting first mixture is adjusted to be acidic. Hydration and fibrosis are then performed sequentially to obtain the protein amyloid fibers. In this invention, the protein powder preferably includes one or more of soy protein isolate (SPI), pea protein isolate (PPI), rice protein (RP), and wheat protein (GP). In this invention, the mass ratio of the protein powder to water is preferably 1:9 to 11, more preferably 1:10. In this invention, the temperature of the first mixing is preferably room temperature, i.e., neither additional heating nor cooling is required, and the time is preferably 10 to 14 hours, more preferably 12 hours. The first mixing is preferably carried out under magnetic stirring. In this invention, the pH value of the acidic mixture is preferably 1.5 to 2.5, more preferably 2. The reagent used to adjust the first mixture to be acidic is preferably a hydrochloric acid solution, and the concentration of the hydrochloric acid solution is preferably 6 mol / L. In this invention, the hydration temperature is preferably room temperature, and the time is preferably 20-35 min, more preferably 30 min. Hydration is preferably carried out under magnetic stirring. After hydration, the invention preferably further includes centrifugation, and the supernatant is collected for fiberization. In this invention, the centrifugation speed is preferably 10000 g, the temperature is preferably 4℃, and the time is preferably 30 min. In this invention, the fiberization temperature is preferably 80-90℃, more preferably 80-90℃, more preferably 85℃; the time is preferably 6-20 h, specifically preferably 6 h, 8 h, 10 h, 12 h, 14 h, 16 h, 18 h, or 20 h. After fiberization, the invention preferably further includes: natural cooling to room temperature and freeze-drying to obtain the amyloid fibers. In this invention, the amyloid fibers are preferably used in the form of an amyloid fiber dispersion, and the concentration of the amyloid fiber dispersion is preferably 25-35 mg / mL.

[0020] This invention also provides a method for preparing the protein amyloid fiber-pyrazine aroma compound described in the above technical solution, comprising the following steps: Amyloid fibers were dispersed in a buffer solution, and then pyrazine aroma compounds were added. The resulting mixture was then equilibrated to obtain the amyloid fiber-pyrazine aroma compound composite material.

[0021] In this invention, the buffer solution is preferably a phosphate buffer solution, the concentration of which is preferably 10 mmol / L, and the pH value is preferably 6.5 to 7.5, specifically 7.2.

[0022] In this invention, the mass concentration of amyloid fibers in the mixture is preferably 0.05-0.15%, more preferably 0.1%, and the concentration of pyrazine aroma compounds is preferably 0.05-0.10 mmol / L, specifically preferably 0.05 mmol / L, 0.06 mmol / L, 0.07 mmol / L, 0.08 mmol / L, 0.09 mmol / L, or 0.1 mmol / L. In this invention, the equilibration temperature is preferably 37°C, the equilibration time is preferably 15-20 h, more preferably 16 h, and the equilibration is preferably carried out under light-protected conditions.

[0023] After balancing, the present invention preferably further includes: freeze-drying the obtained balanced system to obtain the protein amyloid fiber-pyrazine aroma compound.

[0024] The present invention also provides the application of the protein amyloid fiber-pyrazine aroma compound material described in the above technical solution or the protein amyloid fiber-pyrazine aroma compound material prepared by the preparation method described in the above technical solution in plant-based meat products.

[0025] The present invention does not specifically limit the application of the protein amyloid fiber-pyrazine aroma compound, and those skilled in the art can set it according to actual needs.

[0026] The following detailed description, in conjunction with embodiments, illustrates the protein amyloid fiber-pyrazine aroma compound, its preparation method, and its application, but these should not be construed as limiting the scope of protection of this invention.

[0027] Materials and reagents SPI (80% purity) and PPI (80% purity) were purchased from Shanghai Yuanye Biotechnology Co., Ltd. RP (80% purity) was purchased from Shaanxi Mixian'er Biotechnology Co., Ltd. GP (85% purity) was purchased from Anhui Biluchun Biotechnology Co., Ltd. 2-Methylpyrazine (98% purity), 2,5-dimethylpyrazine (98% purity), and 2,3,5-trimethylpyrazine (98% purity) were purchased from Shanghai Maclean's Biochemical Technology Co., Ltd. All other chemicals used were analytical grade, and all water was deionized.

[0028] Example 1: Study using the method of binding 2,5-dimethylpyrazine to protein amyloid fibrils 1. Preparation of amyloid fibers from different protein sources Four different protein powders (SPI, PPI, RP, and GP) were dissolved in 10 times their volume of deionized water and stirred at room temperature for 12 hours. The pH of the solution was adjusted to 2.0 with 6 mol / L hydrochloric acid, and the mixture was magnetically stirred for 30 minutes and centrifuged for 30 minutes (4°C, 10000g). The supernatant was collected and stirred in an 85°C water bath for 20 hours. The mixture was then immediately cooled to room temperature to obtain SAFs, PAFs, RAFs, and GAFs, which were stored at 4°C for later use.

[0029] 2. Test Characterization 2.1 Characterization of amyloid fibrils from different protein sources 2.1.1 Microscopic morphology The concentration of the amyloid fibrous solution was diluted to 0.5 mg / mL and placed on a carbon-coated copper grid. After 5 min, excess sample was removed with filter paper. The sample was stained with phosphotungstic acid, dried, and the microstructure of the amyloid fibrous fibers was observed using a transmission electron microscope (Hitachi HT7700, Japan). The aspect ratio was analyzed using Nano Measurer 1.2 software.

[0030] 2.1.2 Structural Features (1) Thioflavin T 50 μL of amyloid fibrous solution (1 mg / mL) was mixed with 5 mL of thioflavone T staining solution and reacted in the dark for 1 min. The fluorescence curves were collected on a fluorescence spectrometer (FL6500, PerkinElmer, USA) with an excitation wavelength of 440 nm and an emission wavelength of 490 nm.

[0031] (2) X-ray diffraction Freeze-dried amyloid fibers were placed on the current-carrying plate of an X-ray diffractometer (XRD, Smart Lab SE, Rigaku, Japan). The scanning range was 5–90° (2θ), the scanning speed was 5° / min, the voltage was 40 kV, and the current was 40 mA. Diffraction patterns were obtained under Cu-Ka radiation (λ = 0.154 nm).

[0032] (3) Protein spatial conformation: FTIR, fluorescence, ultraviolet Freeze-dried amyloid fibers were ground with spectroscopically pure potassium bromide at a mass ratio of 1:100. FTIR spectra were collected using a Fourier transform infrared spectroscopy (FTIR, Vertex 70, Brooke, Germany) with potassium bromide as background, and 32 scans were performed. The spectral range was 4000–400 cm⁻¹. -1 The resolution is 4cm. -1 The amide I band (1700-1600 cm⁻¹) in the FTIR spectrum was analyzed using PeakFit v4.12 software. -1 The model is fitted and the protein secondary structure is calculated.

[0033] The concentration of the amyloid fibrous solution was diluted to 0.1 mg / mL, and the microenvironmental changes of tyrosine (Δλ=λex-λem=15 nm) and tryptophan (Δλ=λex-λem=60 nm) in the solution were detected using a fluorescence spectrophotometer (FL 6500, PerkinElmer, USA). Excitation wavelengths were 200-400 nm, with wavelength intervals (Δλ) of 15 nm or 60 nm, an excitation slit of 5 nm, and an emission slit of 5 nm.

[0034] The concentration of the amyloid fibrillary solution was diluted to 0.1 mg / mL, and the ultraviolet absorption spectrum was detected using a UV spectrophotometer (UV-2600i, Shimadzu, Japan) with wavelengths of 220-400 nm and data intervals of 0.2 nm. The second derivative plot was obtained using Origin 2024 software.

[0035] (4) Thiol group Total thiol content was determined by adding 15 mg of amyloid cellulose solution to 5 mL of Tris-Gly-8M Urea buffer, and free thiol content was determined by adding 15 mg of amyloid cellulose to 5 mL of Tris-Gly buffer (pH 8.0). Then, 50 μL of DTNB solution (4 mg / mL) was added, and the mixture was incubated at 25°C for 1 h. After centrifugation (25°C, 13600 g, 10 min), the supernatant was collected and measured colorimetrically at 412 nm. The formula for calculating thiol content is as follows: 1.1 SH (μmol / g) = 73.53 × A 412 ×D / C Formula 1.1; In the formula, A 412 The absorbance values ​​are after subtracting the blank. D is the dilution factor, and C is the protein concentration (mg / mL).

[0036] (5) Hydrophobic groups Different concentrations (0.005, 0.010, 0.015, 0.020, 0.025, and 0.030 mg / mL) of protein amyloid cellulose solutions were prepared using K₂HPO₄-KH₂PO₄ buffer (0.01 mol / L, pH 7.2). 4 mL of each solution was added to 20 μL of 8 mmol / L 8-aniline-1-naphthalenesulfonic acid solution, and the solutions were incubated at 25°C in the dark for 5 min before fluorescence intensity was measured. Fluorescence measurement conditions: excitation wavelength 390 nm, emission wavelength 470 nm, emission spectral range 430-520 nm, slit width 1 nm, and graduation 0.5 nm. A linear regression was performed with protein concentration as the independent variable and fluorescence intensity as the dependent variable; the slope represents the surface hydrophobicity.

[0037] (6) Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) Amyloid fibrous solution (0.5 mg / mL) was mixed with loading buffer at a 1:1 ratio, heated in a 100°C water bath for 3 min, cooled to room temperature, and then injected into the sample wells for SDS-PAGE electrophoresis. At the end of electrophoresis, Coomassie Brilliant Blue staining solution was added and stained on a shaker for 12 h, followed by washing with ultrapure water to destain until the electrophoretic bands were clearly visible. Electrophoretic images were collected using a gel imaging system (Bio-Rad Universal Hood II, USA).

[0038] 2.1.3 Physicochemical properties (1) Zeta potential A protein amyloid fibrous sample with a concentration of 0.5 mg / mL was prepared and centrifuged at 3500 r / min for 5 min. The supernatant was collected. The zeta potential of the sample was measured using a Malvern Panalytical Limited (ZSU3100, UK) nanolaser particle size analyzer.

[0039] (2) Particle size and dispersion stability The particle size and polydispersity index of protein amyloid fibers (concentration 0.1 mg / mL) were determined using a nanolaser particle size analyzer (ZSU 3100, Malvern Panalytica science, UK).

[0040] (3) Turbidity The amyloid fibrillation stock solution was diluted 500 times with deionized water and vortexed for 2 min. The absorbance at 600 nm was then measured on a microplate reader (VICTOR Nivo HH3500500, PerkinElmer, USA).

[0041] (4) Solubility The protein amyloid cellulose stock solution was diluted 500 times with deionized water, centrifuged for 15 min (4℃, 10000g), and the protein concentration of the supernatant was measured using a Bradford protein concentration assay kit (Solarbio, Beijing, China) and a microplate reader (VICTOR NivoHH3500500, PerkinElmer, USA).

[0042] (5) Thermal stability The freeze-dried amyloid cellulose was heated from 30°C to 600°C at a rate of 10°C / min using a thermal analyzer (TGA 5500, TA Instruments, USA) to test its thermal stability.

[0043] (6) Rheological behavior Rheological profiles of amyloid filaments were recorded using a hybrid rheometer (Discovery HR-1, TA Instruments, USA). Flow scans were measured at shear rates of 0.1–100 1 / s and at room temperature; frequency scans were measured at a strain of 0.5%, shear rates of 0.1–100 rad / s, and at room temperature.

[0044] (7) Gel network The amyloid fibrous sample was freeze-dried and gold-plated for 60 seconds. The gel network structure of the gel sample at 100 magnification was observed using a scanning electron microscope (HITACHI, SEMS-4800, Japan).

[0045] 2.2 Preparation of protein amyloid fibrils-aroma encapsulation complex Amyloid fibrous samples were dispersed in phosphate buffer (10 mmol / L, pH 7.2), and aroma substance stock solutions were prepared by dissolving 2,5-dimethylpyrazine in chromatographic grade methanol. Appropriate amounts of the amyloid fibrous fibrous solution were mixed thoroughly with the aroma substance stock solutions to achieve a final concentration of 0.1 w / v for the amyloid fibrous ...

[0046] 2.2.1 Determination of aroma binding rate Aroma binding was measured using headspace solid-phase microextraction-gas chromatography (HS-SPME-GC). The aroma-encapsulated complex solution was placed in a headspace vial, sealed with a PTFE-silicone diaphragm, and equilibrated in a 37°C water bath for 10 min. Aroma compounds were extracted and enriched using Carboxen / poly-dimethylsilox fiber (50 / 30µm, 2cm, DVB / CAR / PDMS). The aroma was then analyzed using a gas chromatograph (GC-2010Plus, SHIMADZU, Japan) equipped with a Wax capillary column (30mm × 0.25mm × 0.25μm).

[0047] 2.2.2 Evaluation of aroma binding thermal stability The aroma-encapsulating complex was heat-treated at 50℃ (water bath), 80℃ (water bath), and 120℃ (oil bath) for 10 min, and then equilibrated at 37℃ in the dark for 16 h. The aroma binding rate was determined using the HS-SPME-GC method to evaluate the aroma binding thermal stability under different heat treatments.

[0048] 3. Results and Analysis 3.1 Microscopic morphology of amyloid fibrils from different protein sources Aggregates formed from proteins of different sources after acid-heat treatment all exhibit a linear morphology. Figure 1 (A) demonstrates that acid-heat treatment can induce the transformation of native proteins into fibrous structures. This process may involve two stages: heating under acidic conditions catalyzes the hydrolysis of peptide bonds, breaking the protein down into smaller peptides or free amino acids; subsequently, driven by electrostatic, hydrophobic, and other interactions, the peptides undergo directional aggregation and assembly, ultimately forming specific fibrous structures. Notably, the size distribution of amyloid fibers from different protein sources exhibits differences (…). Figure 2 SAFs exhibit a flexible, linear structure (length: 225.17 nm, diameter: 17.20 nm, aspect ratio: 13.32); PAFs have a similar morphology to SAFs, but are more slender (length: 395.96 nm, diameter: 13.64 nm, aspect ratio: 30.18); RAFs are more worm-like in shape and are generally shorter (length: 63.39 nm, diameter: 13.77 nm, aspect ratio: 4.93); GAFs are unbranched, linear (length: 380.5 nm, diameter: 18.62 nm, aspect ratio: 20.78). This indicates that the morphology of amyloid fibers prepared from natural globulins is closely related to the protein source. In fact, even different components from the same protein source (such as the 7S and 11S components of soybean and pea proteins) form fibers with different morphologies under the same induction conditions. This difference may stem from the variations in the number and conservation of core peptides involved in fibrous assembly in different globulins, thus affecting their final assembly pathway and morphological characteristics.

[0049] 3.2 Structural characteristics of amyloid fibrils from different protein sources 3.2.1 Cross-β-fold structure All four types of amyloid fibrils showed higher thioflavin T fluorescence signals at 488 nm than the original protein. Figure 1 The specific binding of thioflavin T to the cross-β-sheet structure in B) is the reason for the fluorescence surge. This result confirms that all proteins successfully formed amyloid filaments with typical cross-β-sheets after acid-heat treatment. Under the same acid-heat induction conditions, the fluorescence intensity of amyloid filaments formed by legume proteins (SPI and PPI) was significantly higher than that of cereal proteins (RP and GP). This indicates that legume proteins are more conducive to forming cross-β-sheet structures, exhibiting a more efficient amyloid filament formation capacity than cereal proteins under specific induction conditions. This may be due to the different dominant amino acid types of the proteins and the differences in the sensitivity of peptides to acid-heat hydrolysis. Legume globulins are rich in hydrophobic amino acids, which are more easily unfolded and efficiently assembled into cross-β-sheet stacked structures under acid-heat conditions.

[0050] 3.2.2 Crystal Structure All four protein-derived amyloid fibrils exhibited characteristic crystalline diffraction peaks near 2θ=9° (corresponding to the crossed β-structure perpendicular to the protofibril axis, with a slice spacing of 9.82 Å) and 2θ=20° (corresponding to the β-sheet polypeptide backbone parallel to the protofibril axis, with a slice spacing of 4.44 Å). Figure 1 (C) In terms of peak intensity, the amyloid filaments formed by cereal proteins (RP and GP) were significantly higher than those formed by legume proteins (SPI and PPI), indicating that the spatial arrangement of the β-sheet structure in cereal protein fibers may have a higher overall consistency and regularity. This difference may stem from the different molecular compositions and dominant intermolecular forces of the two types of proteins: legume globulins are rich in hydrophobic amino acids, and hydrophobic interactions easily lead to lateral disordered aggregation of β-sheets, thus averaging and weakening the diffraction intensity; while cereal proteins contain more polar amino acids, and their fiber assembly process is more dominated by directional and specific hydrogen bonds, thus resulting in a stronger diffraction signal. It is worth noting that SAFs and PAFs show additional diffraction peaks near 2θ=31.6° and 2θ=45.3°, indicating that the amyloid filaments formed by legume proteins have a higher-order ordered structure, while the fiber structure of cereal proteins is relatively simple, only possessing the core cross-β-sheet feature of amyloid filaments.

[0051] 3.2.3 Protein Spatial Conformation Amide I region (1600-1700 cm⁻¹) in FTIR spectra -1 The absorption peak intensity is mainly affected by the tensile vibrations of C=O and CN. Figure 1The term "D" in this region is often used to reflect changes in protein secondary structure. By performing multi-peak fitting on this region, the relative content of protein secondary structures can be further calculated, and the results are shown in Table 1.

[0052] Table 1. Relative content of protein secondary structures in SAFs, PAFs, RAFs and GAFs solutions.

[0053] Note: Different letters (ad) in the same row indicate significant differences. p <0.05).

[0054] Overall, the four types of amyloid fibers showed low contents of α-helical and random coil structures, while the contents of β-sheet and β-turn structures were relatively high. Notably, legume amyloid fibers (SAFs and PAFs) exhibited high β-sheet content and low β-turn content, while cereal amyloid fibers (RAFs and GAFs) showed the opposite characteristics. This is consistent with previous studies on the secondary structure composition of amyloid fibers induced by soybean and wheat proteins.

[0055] Specifically, there are also significant differences in the same conformation among amyloid fibrils from different protein sources: the proportion of α-helices in GAFs reached 18.25%, which is significantly higher than in other samples ( p<0.05 Compared to other protein amyloid fibers, PAFs have a higher percentage of β-sheets (35.86%). p<0.05 This indicates that, under the same induction conditions, the primary structure of a protein remains the key factor determining the final secondary conformation of amyloid filaments. The inherent amino acid sequence and composition of each protein source determine the characteristics of its core peptide segments for amyloid filament formation, thereby regulating intermolecular interactions during the fibrillation process (such as hydrophobic stacking, hydrogen bond networks, and disulfide bond constraints). Ultimately, this guides proteins through different assembly pathways, forming polymorphic amyloid filaments with cross-beta-sheet conformations but diverse secondary conformations.

[0056] Previous studies have shown that the position of the synchronous fluorescence peak is related to the microenvironment surrounding tyrosine and tryptophan, and a blue shift in the peak usually indicates decreased polarity and increased hydrophobicity. Figure 1 E (Δλ=15nm) and Figure 1 As can be seen from F (Δλ=60nm), under the same acid-heat conditions, RAFs exhibit higher fluorescence intensity, indicating that their tyrosine and tryptophan residues are more encapsulated in the hydrophobic core region. At Δλ=15nm, the positions of the fluorescence peaks of different protein amyloid fibrils are not entirely the same. The maximum emission wavelength of RAFs (λ...) is... maxThe wavelength of Δλ is 280.1 nm, while SAFs (281.6 nm), PAFs (282.7 nm), and GAFs (281.1 nm) all exhibit a red shift to some extent. This difference in shift indicates varying degrees of exposure of tyrosine residues in the protein sequence to the external polar microenvironment. When Δλ = 60 nm, the λ values ​​of SAFs, RAFs, and GAFs are significantly different. max The peak value of PAFs (278.1 nm) was similar (279.6 nm), but showed a blue shift, indicating that the microenvironment of tryptophan residues in PAFs was the least polar. The differences in these fluorescence properties mainly stem from the different numbers, positions, and spatial distributions of aromatic amino acid residues in the fiber structure of different proteins. Therefore, under the same acid-heat treatment conditions, the unique aromatic amino acid sequences of each protein source and their localization in fiber assembly collectively determine the polarity characteristics of the local microenvironment of the residues.

[0057] Ultraviolet second derivative mapping can be used to analyze conformational changes in the microenvironment of aromatic amino acids (tyrosine and tryptophan). Figure 1 (G in the text). All four types of protein amyloid fibrils exhibited positive absorption peaks near 288 nm and 296 nm. The absorption peak at 288 nm was contributed by both tyrosine and tryptophan residues, while the absorption peak at 296 nm represented only the state of tryptophan residues. Since tyrosine is more sensitive to changes in microenvironment polarity, the ratio of the difference between positive and negative absorption peaks in the UV second derivative spectrum (r=a / b) can reflect changes in its surrounding dielectric environment. Studies have shown that an increase in the r value may be related to the unfolding of the protein's tertiary structure and the migration of tyrosine residues to hydrophobic regions. In this example, the r value (2.38) of RAFs was the highest among the four types of protein amyloid fibrils, indicating that the microenvironment polarity of its tyrosine residues was the lowest. This result is consistent with the analysis conclusions of the aforementioned synchronous fluorescence spectroscopy.

[0058] 3.2.4 Thiol group Total sulfhydryl groups are composed of sulfhydryl groups both inside and outside the protein molecule, and their content is related to the number of cysteine ​​residues in the protein sequence. The total sulfhydryl content of RAFs is significantly higher than that of other protein amyloid fibrils (RAFs). Figure 3 A in the middle; p<0.05The total thiol content of SAFs and GAFs is relatively low, indicating a fundamental difference in the number of cysteine ​​residues among different plant amyloid fibrils. During amyloid fibril formation, the protein structure undergoes dramatic rearrangement, and some free thiols may be oxidized to form intramolecular or intermolecular disulfide bonds. Studies suggest that disulfide bonds can positively influence protein conformational stability by reducing the number of unfolded conformations. Therefore, the ratio of free thiols to total thiols can serve as an indicator of the degree of protein unfolding and the strength of intermolecular crosslinking: a lower ratio generally indicates that more free thiols are involved in disulfide bond formation, and the structure may be more compact. In this embodiment, SAFs (63.15%) and RAFs (63.13%) have similar and relatively low ratios of free thiol to total thiol, indicating that more free thiol is oxidized to disulfide bonds during fiber assembly, and the fiber structure may be more compact; while the higher ratios of PAFs (68.28%) and GAFs (66.17%) mean that more free thiol is retained in their fiber structure, and the overall degree of crosslinking is relatively weak.

[0059] 3.2.5 Hydrophobic groups Surface hydrophobicity is often used to describe the contact between hydrophobic groups on a protein surface and a polar aquatic environment. Figure 3 As shown in B, the surface hydrophobicity of PAFs is significantly higher than that of other protein amyloid fibers ( p<0.05 This is mainly attributed to the differences in the primary structure (amino acid sequence) of proteins from different sources. Pea protein, rich in nonpolar hydrophobic amino acids such as leucine and valine, exhibits a higher degree of exposure of its hydrophobic groups under acid-heat-induced fibrosis conditions. Studies have shown that hydrophobic residues on the 11S legumin subunit sequence of pea protein are more likely to remain on the surface after fibrosis, while other proteins (such as chickpeas and lentils) may re-embed hydrophobic regions during fiber assembly. Interestingly, SAFs and RAFs exhibit similar surface hydrophobicity (…). p>0.05 This suggests that despite their different initial amino acid sequences, soybean and rice proteins may have formed similar conformations during fibrosis, resulting in a similar number and distribution of exposed hydrophobic groups. This implies that the formation of protein amyloid fibers may involve a convergent assembly pathway, allowing the final product to transcend differences in the protein's primary sequence in terms of surface properties.

[0060] 3.2.6 SDS-PAGE electrophoresis SDS-PAGE technology can analyze the subunit composition of proteins after fibrillation. SAFs, PAFs, and RAFs all formed small molecular weight protein subunit bands in the region below 11 kDa, but the bands of RAFs in this region were significantly lighter than those of SAFs and PAFs. Figure 3(C) Notably, the electrophoretic bands of wheat protein after fibrillation are concentrated in the low molecular weight range below 5 kDa and the high molecular weight range of 48-75 kDa. This suggests that wheat protein may undergo specific degradation or polymerization during fibrillation: some proteins are hydrolyzed into small peptides, while the remaining subunits form medium-sized aggregates through hydrophobic interactions or covalent cross-linking. Studies have shown that gliadin is easily hydrolyzed into small peptides, which can serve as components in the formation of amyloid fibrils; however, the 10-35 kDa bands gradually decrease with prolonged heating time, indicating that glutenin and gliadin within them gradually participate in the assembly of higher molecular weight aggregates.

[0061] 3.3. Physicochemical properties of amyloid fibers from different protein sources 3.3.1 Zeta potential Zeta potential represents the net charge on the surface of a protein solution. Amyloid filaments formed from different plant proteins all carry a positive charge under acidic conditions. Figure 4 (A) This is because during fibrosis, the solution pH is approximately 2.0, far below the protein's isoelectric point. At this point, the dissociation of the protein's carboxyl groups is inhibited, and the corresponding amides are protonated, resulting in a positively charged solution. However, the absolute value of the zeta potential of PAFs is significantly higher than that of SAFs, RAFs, and GAFs ( p <0.05). This may be because pea protein is rich in basic amino acids such as lysine and arginine, which expose a stronger positive charge density on the surface of amyloid fibers.

[0062] 3.3.2 Particle size, turbidity, and dispersion stability PAFs have a significantly larger particle size than other protein amyloid fibers. p<0.05 ; Figure 4 The presence of B indicates that these fibers are more prone to aggregation or have stronger inter-fiber interactions during fibrosis. Stronger hydrophobic interactions may directly promote the formation of larger aggregates, while potential hydrogen bonds and van der Waals interactions between the β-sheet layers in PAFs may also drive lateral fiber bonding, further increasing particle size. Studies have found a generally positive correlation between particle size and turbidity in natural proteins, and similar trends are observed in the turbidity and particle size of amyloid fibers from different plant protein sources. However, it is noteworthy that RAFs and GAFs exhibit significantly different particle sizes (…). p<0.05 ) and similar turbidity ( p>0.05 This reflects that the turbidity of amyloid fibers is not entirely determined by particle size, but is also related to other factors such as particle shape and aggregation state. Dispersion stability reflects the width of the particle size distribution in the sample; generally, the lower the dispersion stability, the more uniform the particle size distribution. GAFs have the smallest particle size and the most stable dispersibility. Figure 4The C in this context is due to the smaller particle size resulting in a slower settling rate, which enhances the physical stability of the dispersion.

[0063] 3.3.3 Solubility Protein solubility is the outward manifestation of the thermodynamic equilibrium reached between protein-protein and protein-solvent interactions, and its changes are determined by both average hydrophobicity and net surface charge: hydrophobic interactions promote protein-protein aggregation, reducing solubility; ion hydration enhances protein-water affinity, aiding dissolution. After acid-heat treatment, SAFs and PAFs have high absolute values ​​of Zeta potential (net surface charge characteristics), and theoretically, their solubility can be improved through ion hydration. However... Figure 4 The results from D show that SAFs and PAFs, both belonging to the legume protein family, have similar and low solubilities. This indicates that hydrophobic interactions dominate the inhibition of solubility, with protein-protein interactions being stronger than protein-water interactions, thereby driving fiber aggregation and inhibiting dissolution. In contrast, RAFs have significantly higher solubility than other protein amyloid fibers (...). p< 0.05 This may be because the strong surface charge in RAFs solution enhances the electrostatic repulsion between protein molecules while promoting ion hydration, attracting more water molecules to bind to the protein surface. This enhanced protein-water interaction effectively counteracts the aggregation tendency caused by hydrophobic interactions, thereby improving the water solubility of RAFs.

[0064] 3.3.4 Thermal stability The thermal degradation of different plant protein amyloid fibers can be divided into three stages ( Figure 4The initial stage (below approximately 150℃) of mass loss is mainly attributed to the evaporation of residual moisture. The maximum thermal degradation temperature of RAFs in this stage is 75.72℃, significantly higher than that of SAFs (58.86℃), PAFs (60.48℃), and GAFs (63.10℃), indicating that RAFs have a strong water retention capacity. The second stage (150-400℃) of mass loss is mainly caused by the thermal degradation of the protein structural backbone. In this stage, the maximum thermal degradation temperatures of different amyloid fibers are 323.66℃ (SAFs), 320.89℃ (PAFs), 312.92℃ (RAFs), and 318.72℃ (GAFs), respectively. This shows that legume amyloid fibers exhibit stronger thermal stability in high-temperature environments compared to cereal amyloid fibers. Studies have shown that the thermal stability of natural amyloid fibers is related to the content of β-sheet structures: β-sheets form a stable "three-dimensional zipper" structure through main-chain hydrogen bonds and hydrophobic interactions and van der Waals forces between side chains, thereby enhancing the thermal stability of amyloid fibers. The higher proportion of β-sheet conformations in legume amyloid fibers may be the structural basis for their greater resistance to degradation at high temperatures.

[0065] 3.3.5 Rheological Behavior The rheological behavior of proteins is closely related to their gel quality, with storage modulus (G', representing solid behavior) and loss modulus (G'', representing liquid behavior) being important indicators of gel viscoelasticity. Frequency scanning revealed that, at the same angular frequency, the storage modulus (G') and loss modulus (G'') of PAFs and SAFs were higher than those of RAFs and GAFs. Figure 4 The FG in the figure indicates that the fiber network formed by legume protein amyloid fibers has stronger viscoelasticity. This may be due to the high β-sheet ratio in legume protein fibers, which promotes inter-fiber cross-linking through hydrogen bonding and hydrophobic interactions, forming a denser three-dimensional network. All protein amyloid fibers exhibit shear-thinning behavior characteristic of pseudoplastic fluids, that is, the apparent viscosity decreases with increasing shear rate. Figure 4 The H in this figure is due to the shearing action disrupting the entanglement and connection between molecular chains. At the same shear rate, the apparent viscosity of legume amyloid fibers is greater than that of cereal amyloid fibers. This may be because legume amyloid fibers have a denser spatial network structure, with tighter entanglement between molecular chains, resulting in greater resistance to fluid flow.

[0066] 3.3.6 Microstructure The microstructures of different plant protein amyloid fibers show significant differences. Figure 4In the diagram (I): SAFs exhibit a dense and irregular sheet-like structure. PAFs have a relatively large sheet-like gel structure with a smooth surface. RAFs present as fragmented sheets and short bundles of fibers, with a blurred and disordered sheet structure. GAFs are porous sheets with many protrusions. These differences in microscopic morphology indicate that their self-assembly pathways are polymorphic due to differences in protein amino acid composition and physicochemical properties (hydrophobicity, charge, hydrolysis sensitivity, etc.). Even for the same protein, different environmental conditions (such as pH, ionic strength, temperature, and heating time) can profoundly affect its aggregation pathway and final morphology. For example, β-lactoglobulin forms typical amyloid filaments at pH 2, while at pH 3.5 it mainly forms worm-like amorphous aggregates. This difference can be attributed to the effect of pH on the degree of acid hydrolysis: at lower pH, proteins are more easily hydrolyzed into peptides, which become the basic units for fiber assembly; while at pH conditions close to the isoelectric point, the overall protein structure is preserved more, tending to form disordered aggregates.

[0067] 3.4 Aroma-binding capacity of amyloid fibers from different protein sources 3.4.1 Aroma Binding Rate SPME-GC technology was used to evaluate the differences in the ability of amyloid fibers from different protein sources to adsorb aroma compounds. For example... Figure 5 As shown in result A, the binding rates of the four protein amyloid fibrils to 2,5-dimethylpyrazine were significantly higher than those to the original protein. p<0.05 This indicates that modifying the fibrous structure of proteins is an effective way to enhance their flavor-binding properties. This may be because when natural proteins are converted into amyloid fibers by acid and hot water hydrolysis, the proteins dissociate from their native folded state, and the peptide chains reorganize into a cross-beta-sheet structure with hydrogen bond networks and hydrophobic side chain stacks as the core, exposing more hydrophobic regions or forming specific binding pockets, thereby more effectively binding 2,5-dimethylpyrazine.

[0068] The binding rates of four protein amyloid fibers to 2,5-dimethylpyrazine were 36.64% (PAFs), 23.73% (RAFs), 23.49% (SAFs), and 16.15% (GAFs), respectively, indicating that the adsorption capacity of amyloid fibers from different protein sources for flavor compounds is specific. First, protein amyloid fibers rich in aromatic or hydrophobic residues readily bind to hydrophobic aromatic heterocyclic compounds through hydrophobic interactions and π-π stacking. Pea protein typically contains a high proportion of hydrophobic and aromatic amino acids; during acid-thermal hydrolysis-induced fibrillation assembly, these residues tend to be exposed on the fiber surface, forming abundant hydrophobic microdomains and aromatic ring arrays, thus providing numerous binding sites for 2,5-dimethylpyrazine. Second, the morphology and aggregation structure of different protein amyloid fibers directly affect their physical retention efficiency for aroma molecules. PAFs have a high aspect ratio ( Figure 2 ) and a regular, ordered fiber network structure ( Figure 4 The I in PAFs provides a larger effective specific surface area and more binding sites, which further increases the accessibility of aroma molecules. In addition, the electrostatic microenvironment on the surface of PAFs may also affect the adsorption of charged or polar aroma molecules. Studies have shown that electrostatic interactions are an important adsorption mechanism. PAFs have a strong positive charge density (… Figure 4 A) in the above may readily generate electrostatic attraction with polar groups in aroma molecules (such as nitrogen atoms on pyrazine rings), further enhancing its affinity for pyrazine flavor compounds.

[0069] 3.4.2 Aroma binding thermal stability Overall, the aroma-binding capacity of all protein amyloid fibers decreased with increasing temperature, reaching its lowest point at 120°C. Figure 5 (B in the figure) indicates that high temperature disrupts the binding between protein amyloid fibers and aroma molecules. On one hand, this may be because increased temperature increases the thermal kinetic energy of aroma molecules, making them more likely to desorb from the binding sites on the surface of protein amyloid fibers, thus leading to a decrease in binding rate. On the other hand, extreme high temperatures may cause the already formed amyloid fibers to break or depolymerize, disrupting their regular and ordered network structure, thereby reducing the effective specific surface area and binding sites, leading to aroma release. Specifically, under different temperature conditions, PAFs can maintain a high aroma binding capacity ( Figure 5 The presence of C in the text reflects the relatively good thermal stability of its aroma complex. PAFs possess a high content of cross-beta-sheet core structures, which can resist certain thermal disturbances during the heating process from 37℃ to 120℃, and are not easily depolymerized or denatured due to temperature increases. This allows the overall framework of PAFs to remain relatively intact after heat treatment, providing a stable "scaffold" for the binding of aroma molecules.

[0070] Example 2: A method utilizing soybean protein starch-like fibers to bind pyrazine aroma compounds with varying numbers of methyl groups. 1. Preparation of SAFs with different acid heat treatment times SPI powder was completely dissolved in 10 times its volume (w / v) of deionized water. The pH of the solution was adjusted to 2.0 with 6 mol / L HCl, and the solution was magnetically stirred for 30 min before centrifugation (4℃, 10000g, 30 min). The supernatant was collected and divided into three equal portions, which were then heat-treated in an 85℃ water bath for 6 h, 12 h, and 18 h, respectively. The samples were immediately cooled to room temperature after heat treatment, and the resulting soybean protein amyloid fibers (SAFs) were stored at 4℃ for subsequent analysis. The SAFs heat-treated for 6 h, 12 h, and 18 h were named SAFs-6, SAFs-12, and SAFs-18, respectively.

[0071] 2. Test Characterization 2.1 Characterization of SAFs under different acid heat treatment times 2.1.1 The microstructure is the same as 2.1.1 in Example 1.

[0072] 2.1.2 The structural features are the same as those in 2.1.2 of Example 1.

[0073] 2.1.3 The physicochemical properties are the same as those in 2.1.3 of Example 1.

[0074] 2.1.4 Yield of SAFs under different acid heat treatment times SAFs were prepared according to the experimental steps in section 1, with water bath heating times set to 6 h, 12 h, 18 h, and 24 h, respectively. After acid heat treatment, cooling and hydration were performed to form a mature SAF dispersion, which was then freeze-dried to obtain SAF powder, which was accurately weighed. The yield of SAFs was defined as the percentage of the mass of the freeze-dried SAF powder to the mass of the initially added SPI powder, calculated using formula 2.1 as follows: Formula 2.1; In Formula 2.1: m SAFs Mass (g) of SAFs powder obtained after freeze-drying; m SPI : Initial mass of SPI powder (g).

[0075] 2.2 Preparation of SAFs-pyrazine aroma complexes SAFs samples were dispersed in phosphate buffer (10 mmol / L, pH 7.2). 2-methylpyrazine, 2,5-dimethylpyrazine, and 2,3,5-trimethylpyrazine were dissolved in chromatographic grade methanol to prepare aroma substance stock solutions (60 mmol / L). Appropriate amounts of SAFs samples were mixed thoroughly with the different aroma substance stock solutions to achieve a final concentration of 0.1 w / v% for the amyloid fibrils and a final aroma substance concentration of 0.08 mmol / L. The mixed solutions were equilibrated at 37°C in the dark for 16 h to allow for sufficient binding of the SAFs and aroma substances, thus obtaining aroma-encapsulated complexes. The three aroma complexes were named SAFs-M, SAFs-D, and SAFs-T, respectively.

[0076] 2.2.1 The determination of aroma binding rate is the same as in 2.2.1 of Example 1.

[0077] 2.2.2 The evaluation of aroma binding thermal stability is the same as in 2.2.2 of Example 1.

[0078] 3. Results and Discussion 3.1. Microstructure of SAFs Soy protein isolate is mainly composed of globulins, accounting for approximately 80-90%. Heating under acidic conditions (pH 2.0) can induce significant fibrotic morphological changes in it. Figure 6 (A) As the heat treatment time increased, SPI exhibited a typical fibrosis evolution process: after 6 hours of treatment, a large number of strip-shaped particles of varying sizes and uneven distribution appeared in the sample, and some spherical structures began to transform into protein amyloid fibrous structures; after 12 hours of heating, protofibrils gradually aggregated; by 18 hours, protein protofibrils tended to form dense aggregates, with some undergoing excessive aggregation to form linear clusters. The above results indicate that acid heat treatment can promote a time-dependent fibrosis process in soybean protein. This process may originate from the destruction of the protein's secondary structure by acidic heating conditions, leading to denaturation and exposure of hydrophobic regions, followed by protein self-assembly driven by hydrophobic interactions. Further size analysis was performed, and the results are shown in Table 2.

[0079] Table 2. Length, diameter, and aspect ratio of SAF under three different hot acid treatment times.

[0080] Note: Different letters (ac) in the same line indicate significant differences. p <0.05).

[0081] With prolonged acid heat treatment, the average length of amyloid fibers gradually increased (70.58-178.47 nm). p <0.05, the average diameter gradually decreases (10.52-8.59 nm, p<0.05, the aspect ratio increases accordingly (6.82-21.73, p <0.05). The above quantitative data indicate that acid heat treatment not only induces SPI to form fibrous structures but also promotes its maturation, manifested as the gradual elongation, thinning, and densification of protein amyloid fibers. It is evident that acid heat treatment can induce a typical and controllable fibrillatory self-assembly process in proteins.

[0082] 3.2 Structural characteristics of SAFs 3.2.1 Cross-β-fold structure The core structural feature of amyloid fibrils is a highly ordered cross-β-sheet. Thionyl T, a cationic benzothiazole fluorescent dye, can specifically insert into and bind to the cross-β-sheet structure, thereby significantly enhancing the fluorescence intensity. SPI exhibits the lowest thiosulfate T fluorescence intensity at 487 nm; however, after acid heat treatment, the fluorescence intensity of the fibrotic samples increases significantly with treatment time, showing a clear time dependence. Figure 6 (B in the text). This result indicates that acid heat treatment can effectively induce fibrosis of SPI and form protein amyloid fibers rich in cross-beta-sheet structures.

[0083] 3.2.2. Crystal Structure X-ray diffraction patterns showed that all samples exhibited characteristic peaks at 9.00° and 19.18° (corresponding to the interchain spacing of α-helices d≈9.8Å and the interchain spacing of β-lamellae parallel to the fiber axis d≈4.6Å, respectively), consistent with the typical characteristics of the basic crystal structure of proteins. Figure 6 (C in the text). However, compared to SPI, the characteristic peak intensities of SAF samples were all slightly reduced, indicating that the fibrosis process disrupted the crystalline order of proteins. This phenomenon can be attributed to the conformational transition of proteins induced by acid-heat treatment: heat treatment breaks some of the secondary bonds (such as hydrogen bonds) that maintain spatial structure, causing peptide chain dissociation, molecular stretching, and reducing ordered structure while increasing disordered structure, thereby reducing crystallinity. Notably, SAFs exhibited additional diffraction peaks at 31.8° and 45.56°, and the peak values ​​significantly increased with prolonged acid-heat treatment time. These new peaks may originate from novel ordered structures formed during fibrosis, such as parallel or cross-beta-sheet structures.

[12] The peak enhancement indicates that, with prolonged processing time, the β-sheet structure tends to stack in a more ordered manner, reflecting the evolution of the fibrous structure towards a more mature and ordered state.

[0084] 3.2.3. Protein Spatial Conformation Protein secondary structure: Amide A band (3000-3500 cm⁻¹) in FTIR spectrum -1The absorption peak originates from the stretching vibrations of NH and OH, and its intensity is mainly affected by hydrogen bonding. The peak intensity in this region increases with increasing acid heat treatment time for all samples. Figure 6 The D in the figure indicates that the intermolecular hydrogen bonds are strengthened in a time-dependent manner during fibrosis, a change that helps improve the stability of protein conformation and overall order. Amide I region (1600-1700 cm⁻¹) -1 The secondary structures of SPI and SAFs are related to the bending oscillations of C=O and CN in protein secondary structures. Quantitative analysis of the protein secondary structures of SPI and SAFs was performed, and the results are shown in Table 3.

[0085] Table 3 Protein secondary structure content of SPI and SAFs

[0086] Note: Different letters (ac) in the same line indicate significant differences. p <0.05).

[0087] Table 3 shows that the acid-heat treatment-induced fibrosis process significantly increased the relative content of β-sheets, exhibiting a time-dependent increase. Compared with SPI (35.11%), the relative increases in β-sheet content in SAFs-6, SAFs-12, and SAFs-18 were 1.96%, 3.50%, and 5.27%, respectively. p <0.05. Meanwhile, the ratio of α-helices to β-turns decreased significantly, while the proportion of random coils increased ( p <0.05). These changes are consistent with previous studies showing that acid heat treatment drives the transformation of soybean protein from α-helices to β-sheets. This transformation may originate from acid heat treatment inducing partial unfolding of the protein and exposure of hydrophobic groups, which in turn triggers α-helix unwinding and promotes structural rearrangement, ultimately forming an ordered fibrous structure dominated by β-sheets.

[0088] Synchronous fluorescence: Synchronous fluorescence spectroscopy, through scanning with a fixed wavelength difference (Δλ), can effectively detect changes in the chromophore microenvironment: Δλ = 15 nm mainly reflects the characteristics of tyrosine residues, while Δλ = 60 nm corresponds to the characteristics of tryptophan residues. After acid-heat treatment and fiberization, the maximum emission wavelength (λ) of tyrosine residues in SPI... max The wavelength of tryptophan residues shifted from 294.9 nm (SPI) to 284.1 nm (SAFs) due to a blue shift. max The fluorescence intensity shifted from 287.3 nm (SPI) to 284.0 nm (SAFs), and the fluorescence intensity of both gradually decreased. Figure 6 (EF in the text). λ maxThe blue shift indicates a decrease in the polarity and an increase in the hydrophobicity of the microenvironment surrounding tyrosine and tryptophan residues. This may be due to conformational changes induced by acid-heat treatment, causing residues to migrate from hydrophilic environments to hydrophobic regions. The decrease in fluorescence intensity may be related to protein denaturation and structural rearrangement caused by acid-heat treatment. Some proteins form amorphous aggregates during this process, thus obscuring the fluorescence of internal tyrosine and tryptophan residues. It is noteworthy that λ... max The fluorescence intensity had already shifted to the blue position at SAFs-6 and remained stable during subsequent treatments. This indicates that the polarity of the chromophore microenvironment had already undergone a fundamental shift in the early stages of fibrosis. The continued decrease in fluorescence intensity with prolonged treatment time likely reflects further optimization of the length, density, and internal arrangement of the amyloid fiber structure, rather than a sustained change in microenvironmental polarity.

[0089] Second-order ultraviolet (UV) spectroscopy: UV second-order spectroscopy is an effective means of expressing local conformational changes of aromatic amino acid residues. All four samples exhibited two positive absorption peaks near 288 nm and 295 nm, and two negative absorption peaks near 284 nm and 292 nm. Figure 6 The positive absorption peak at 288 nm is attributed to both tyrosine and tryptophan residues, while the positive absorption peak at 295 nm is specifically attributed to tryptophan residues. After fibrillation, both positive absorption peaks exhibit a blue shift, indicating a decrease in the polarity and an increase in the hydrophobicity of the microenvironment surrounding the tryptophan residues. Furthermore, the ratio of the difference between the positive and negative absorption peaks (r = a / b) reflects changes in the dielectric microenvironment surrounding the tyrosine residues. Studies show that the unfolding of the protein's tertiary structure and the entry of tyrosine residues into hydrophobic regions lead to an increase in the r value. It was found that the r value is positively correlated with the acid heat treatment time, gradually increasing from 1.07 (SPI) to 1.47 (SAFs-6), 1.61 (SAFs-12), and 1.78 (SAFs-18). This implies that the higher the degree of fiberization in soybean protein, the more hydrophobic the microenvironment surrounding the tyrosine residues. This is consistent with the conclusions obtained from the synchronous fluorescence spectroscopy analysis above, jointly revealing that the fibrillation process drives aromatic amino acid residues into a more hydrophobic microenvironment.

[0090] 3.2.4. Thiol group Total thiol groups typically reflect the degree of protein denaturation, including both surface-exposed free thiol groups and thiol groups embedded within the molecule. During soybean protein fibrosis, changes in both free and total thiol groups reveal the protein structure remodeling process under acid-heat treatment. Figure 7 (A) In the early stages of fibrosis, the protein conformation has not yet fully unfolded, and the internally embedded thiol groups are not effectively exposed. Although the environment changes, the spatial accessibility of these thiol groups remains low, therefore the content of free thiol groups does not show significant changes (A). p>0.05). Meanwhile, the total thiol content decreased at this stage, possibly because protein molecules formed dense aggregates through non-covalent interactions such as hydrophobic interactions, creating a physical masking effect that made it difficult for the detection reagent to reach and react, thus reducing the apparent total thiol content. With prolonged processing time, acidic and thermal conditions drove the protein to undergo thorough and deep unfolding, gradually opening its higher-order structure and exposing more previously embedded thiol groups to the molecular surface. Simultaneously, thiol groups that were initially masked by aggregation became accessible again due to structural rearrangement. The weakening of the physical masking effect and the increased accessibility of functional groups together contributed to a rebound in the detected total thiol content.

[0091] 3.2.5. Hydrophobic groups Hydrophobic interactions are one of the key driving forces for maintaining the tertiary structure of proteins. In this embodiment, the changes in surface hydrophobicity exhibit obvious stage-like characteristics. Figure 7 (B in the original text). In the early stages of fibrosis, protein molecules partially unfold, and although a certain number of hydrophobic groups migrate from the interior to the surface, the surface hydrophobicity does not increase significantly because the overall structure has not yet fully reorganized into typical amyloid fibers. p >0.05). This phenomenon may be attributed to the fact that although the α-helix unwinding releases some hydrophobic groups, the insoluble aggregates formed by the 11S globulin basic polypeptide and the β subunit of the 7S globulin mask some hydrophobic sites, thus inhibiting a significant increase in surface hydrophobicity. As the fibrosis process progresses, under the synergistic effect of acid and heat, the protein undergoes further hydrolysis and conformational rearrangement, causing more hydrophobic groups originally embedded inside the molecule to be exposed to the hydrophilic environment. Compared with SPI, the surface hydrophobicity of SAFs-12 increased significantly by 105.80%, while that of SAFs-18 increased by 168.14% (…). p <0.05). Furthermore, the observed trends in surface hydrophobicity and α-helix content in this embodiment are consistent with the structure-activity relationship reported in the literature, i.e., α-helix content is significantly negatively correlated with surface hydrophobicity. With the reduction of α-helix structure, the protein molecule conformation tends to be looser, and internal hydrophobic residues are more easily exposed to the surface, thereby promoting an increase in surface hydrophobicity.

[0092] 3.2.6. Gel Network Structure The electrophoresis images show clear protein bands. Figure 7 The protein subunits of C in the formula were analyzed in different molecular weight ranges, and the results are shown in Table 4.

[0093] Table 4. Absolute quantitative ratios of SPI and SAF solution electrophoresis bands

[0094] Note: "—" indicates an absolute ratio that was not detected. The data in the table are the average of three measurements. Different letters (ad) in the same row indicate significant differences. p <0.05).

[0095] Table 4 shows that fibrosis modification significantly altered the subunit distribution of the protein. In the high molecular weight range of 90-45 kDa, only SPI showed an absolute quantitative ratio (target protein gray value / maker protein gray value), while no obvious bands were detected in SAFs within this range, indicating that components in this molecular weight range of natural soybean protein were effectively degraded during acid-heat treatment. In the 35-10 kDa range, the absolute quantitative ratio of SPI was significantly higher than that of the SAFs group, and this ratio in the SAFs group showed a significant decreasing trend with increasing acid-heat treatment time. p The value <0.05 indicates that protein components in this range are gradually degraded or participate in the assembly of amyloid fibers under acid-heat conditions, exhibiting a clear time dependence. In the low molecular weight range of 7-4 kDa, the absolute quantitative ratio of SAFs samples is positively correlated with acid-heat treatment time, indicating that low molecular weight peptides gradually accumulate during this process. The changes in protein components in the above different molecular weight ranges collectively reveal the dynamic process of soybean protein fibrillation: that is, medium and high molecular weight proteins gradually degrade over time, while low molecular weight peptides continuously accumulate and participate in orderly assembly. This trend is basically consistent with the reported process of kidney bean protein fibrillation, further supporting that peptide hydrolysis or the release of smaller peptides are key factors in the formation of amyloid fibers from legume globulins under acidic conditions.

[0096] 3.3 Physicochemical Properties of SAFs 3.3.1. Potential The zeta potential refers to the potential of the shear layer on the surface of charged particles, and is widely used to characterize the stability of protein suspensions and the electrostatic interactions between particles. After fibrillation, the zeta potential of SPI changed from -34.04 mV to 21.89 mV (SAFs-6), and gradually increased to 25.43 mV (SAFs-12) and 30.45 mV (SAFs-18) with increasing acid-heat treatment time. Figure 8 (A) The change in potential from negative to positive is mainly due to the system's pH (approximately 2.0) being much lower than the isoelectric point of soybean protein, causing the surface carboxyl groups (such as the -COO groups of glutamic acid and aspartic acid residues) to change from negative to positive. -The protein is gradually protonated to -COOH, thereby reducing the contribution of negative charge and making the system positively charged. As the treatment time increases, the Zeta potential further increases, which may be related to the formation of protein amyloid filament structure and changes in protein conformation: acid heat treatment promotes partial hydrolysis of protein, reducing negatively charged groups on the surface (such as acidic amino acid residues); at the same time, heating destroys the secondary structure of protein, exposing the originally embedded hydrophobic regions and positively charged amino acid side chains (such as lysine and arginine), which together increase the surface positive charge density.

[0097] 3.3.2 Particle size and turbidity Particle size and turbidity can serve as macroscopic indicators of the size and number of suspended particles in aqueous solutions, reflecting the aggregation state of proteins. Regarding particle size, SPI showed a significant decrease of 25.09% after 6 hours of fibrillation. Figure 8 B in the middle; p <0.05). This change may be related to electrostatic interactions caused by pH changes in the system: SAFs have a pH of approximately 2.0, which is farther from the isoelectric point of proteins (pI between 4.5 and 5.2) compared to SPI (pH approximately 6.5). Stronger electrostatic repulsion may inhibit the thermal aggregation of proteins, leading to a decrease in particle size. However, with prolonged acid-heat treatment, the particle size significantly recovers. p <0.05%, which may be because continuous heating further denatures the protein, exposing more hydrophobic regions and driving the formation of larger aggregates. Previous studies have shown a positive correlation between protein turbidity and particle size changes; low turbidity indicates smaller particles and weaker light scattering, while increased turbidity reflects larger particle size. The results in this example are consistent with this trend: the turbidity of SPI decreased significantly after acid-heat treatment, indicating that protein particles were refined in the early stages of fibrosis; however, with prolonged treatment time, the turbidity of SAFs significantly increased (…). Figure 8 (B in the text). The simultaneous increase in turbidity and particle size of SAFs suggests that after initial fibrosis and dispersion, continuous acid-heat treatment may induce further cross-linking, lateral aggregation, or supramolecular network formation of SAFs, thereby assembling them into large aggregates with stronger light scattering capabilities.

[0098] 3.3.3 Solubility and Dispersion Stability Protein solubility essentially reflects the thermodynamic equilibrium state of protein-protein interactions and protein-solvent interactions in a dispersion system. Hydrophobic interactions promote protein aggregation and reduce solubility, while ion hydration enhances protein-water interactions and increases solubility. In this embodiment, the solubility of SAFs was significantly lower than that of SPI and decreased in a time-dependent manner. p<0.05), among which SAFs-6, SAFs-12 and SAFs-18 decreased by 30.44%, 50.82% and 58.05% respectively. Figure 8 C in the text). The numerous hydrophobic regions exposed during fibrosis significantly enhance hydrophobic interactions (C). Figure 7 In the case of B), its effect on promoting aggregation outweighs the dissolution-promoting effect caused by the increase in surface charge. Figure 8 The presence of A in the polydispersity index ultimately leads to a decrease in solubility. Therefore, during the fibrosis of SPI, hydrophobic interactions play a dominant role in inhibiting dissolution behavior. The polydispersity index can be used to assess the homogeneity and stability of protein dispersion systems; a lower value indicates a more uniform particle distribution and a more stable system. The polydispersity index showed no significant change in SPI after 6 hours of fibrosis. p >0.05), but as the processing time was extended to 12 and 18 hours, the time dependence of this index decreased ( Figure 8 D in the middle; p <0.05). This may be because proteins tend to randomly aggregate after unfolding in the early stages of fibrosis, resulting in a wider particle distribution; while prolonged acid-heat treatment drives aggregate reconstruction, forming a more uniform particle structure through fusion or ordered rearrangement, thereby significantly reducing the polydispersity index.

[0099] 3.3.4 Thermal stability The mass loss of SAFs samples mainly occurred between 150 and 600 °C. During this stage, the maximum thermal degradation temperature increased from 308.66 °C (SPI) to 311.50 °C (SAFs-6), 314.91 °C (SAFs-12), and 315.56 °C (SAFs-18); simultaneously, the mass loss rate decreased from 66.61% (SPI) to 65.56% (SAFs-6), 64.77% (SAFs-12), and 63.82% (SAFs-18). Figure 8 The increase in the maximum thermal degradation temperature (E) reflects enhanced thermal stability, while the decrease in mass loss rate suggests a reduced degree of degradation. This phenomenon may be attributed to the synergistic effect of the strengthening of the hydrogen bond network and hydrophobic interactions during fibrosis. On the one hand, the increased thermal motion of molecular chains during SPI fibrosis leads to the full extension and re-entanglement of the molecular chains, forming a denser intermolecular hydrogen bond network in the β-sheet region. This enhanced hydrogen bonding effectively improves the thermal stability of the material, manifested as a gradual increase in the maximum thermal degradation temperature. On the other hand, the enhanced hydrophobic interactions during fibrosis promote the formation of dense hydrophobic microdomains, which not only increases the molecular stacking density but also reduces the hydration capacity.

[31] This reduces mass loss during thermal decomposition. The synergistic effect of the strengthening of the hydrogen bond network and hydrophobic interactions constitutes the structural basis for the improved thermal stability of SAFs.

[0100] 3.3.5 Rheological Behavior Within the range of 0.1–100 rad / s, the storage modulus (G') and loss modulus (G'') of the four protein samples continuously increased with increasing angular frequency, indicating that a solid-like network structure based on entanglement was formed within the system under dynamic shearing. Figure 8 (FG in the sample). With prolonged heat treatment time, the G' and G'' of each sample showed a time-dependent increase across the entire frequency range. Among them, SPI had the lowest modulus value, indicating that its intermolecular entanglement was loose, and it could only form a primary and unstable elastic network with limited resistance to external deformation. After fibrillation treatment, the G' and G'' of SAFs showed a time-dependent increase. This change indicates that acid heat treatment-induced protein self-assembly effectively strengthened the network structure of the system. From the perspective of structural evolution, acid heat treatment promotes the bonding, breaking, and rearrangement of protein molecules, forming protein amyloid fibers rich in β-sheets. These protein amyloid fibers are further cross-linked through enhanced hydrogen bonding and hydrophobic interactions, constructing a denser and more stable three-dimensional network, thereby significantly improving the mechanical strength of the protein system and making it exhibit stronger elastic and viscous responses under dynamic shear.

[0101] Viscosity is a key rheological parameter characterizing the internal resistance of a fluid. All samples exhibited typical pseudoplastic fluid behavior, meaning that the apparent viscosity decreased significantly with increasing shear rate, demonstrating a clear shear thinning phenomenon. Figure 8 (H in the text). From a microstructural perspective, at low shear rates, proteins form a three-dimensional entangled network through hydrogen bonds and hydrophobic interactions, resulting in high flow resistance. As the shear rate increases, the network structure is disrupted, and molecules align along the flow direction, leading to a decrease in resistance. Notably, the shear thinning degree of the samples significantly increased after 12 hours of fibrillation. This is closely related to the enhanced hydrophobic interactions: hydrophobic groups transition from an "exposed state" to a "cross-linked state," promoting the formation of a denser network and thus increasing the effective volume of the system. The increase in effective volume means that proteins form a more robust network when at rest, but this network is more easily disrupted during flow. This explains why SAFs-12 and SAFs-18 simultaneously possess high initial viscosity and a significant shear thinning effect, demonstrating the dual regulatory role of fibrillation treatment on protein rheological behavior.

[0102] 3.3.6 Scanning Electron Microscopy The microstructures of the four sample gels are shown in the figure ( Figure 8In SAFs-12, the natural SPI exhibits a regular, sheet-like aggregate with the highest structural order. This is due to the precise embedding of the hydrophobic core in its natural conformation, the stability of the hydrogen bond network, and the synergistic interaction of various processes, all of which together constitute the thermodynamically lowest energy state. SAFs-6 show structural fragmentation and dispersion, with only a preliminary, irregular aggregated morphology. This indicates that the protein conformation unfolding under acid-heat treatment is insufficient, and the molecular assembly process has not yet achieved effective coordination. In SAFs-12, the fragments begin to align directionally, forming a more regular network structure locally. The fibrous network of SAFs-18 further tends to be regular, compact, and ordered. These structural evolutions reveal that acid-heat treatment drives proteins through a dynamic "unfolding-reassembly" process. Essentially, this process is the structural evolution of protein molecules under acid-heat conditions far from equilibrium, from the thermodynamically stable natural folded state (SPI), through a kinetically controlled assembly intermediate state (SAFs-6 / 12), and finally into a novel, stable fibrous assembly state (SAFs-18). It is worth noting that the connotation of "structural order" has undergone a fundamental change in this process: from short-range order in the natural state (dependent on specific three-dimensional folding) to long-range order in the state of protein amyloid filaments (based on the extension and stacking of cross-β-sheets).

[0103] 3.4 Yield of SAFs Acid-heat treatment time significantly affected the formation efficiency of SAFs, with the yield initially increasing rapidly and then stabilizing over time. No significant difference in yield was observed between 18 h (30.02%) and 24 h (30.60%). Figure 8 J in the middle; p The result >0.05 indicates that the fibrosis reaction had essentially reached dynamic equilibrium within 18 hours. This result further confirms that only certain specific hydrolyzed polypeptide fragments in the fibrosis process have the ability to assemble into amyloid fibers, and the yield no longer increases linearly with time when these precursors are depleted or reach equilibrium.

[0104] Considering both preparation efficiency and product yield, acid heat treatment for 18 hours was determined to be the optimal process parameter for preparing protein amyloid fibers. Although 24 hours yielded the highest yield, it did not show a significant statistical gain compared to 18 hours and was accompanied by higher time and energy costs. Therefore, to balance high yield with economic efficiency, this embodiment ultimately selected SAFs-18 as the ideal protein carrier for further in-depth investigation of aroma binding rate and interaction mechanisms.

[0105] 3.5. Binding ability of SAFs to pyrazine aroma compounds with different numbers of methyl groups 3.5.1. Aroma Binding Rate The aroma-binding capacity of proteins is closely related to their spatial conformation and the chemical structure of aroma molecules. Current research focuses primarily on natural globulins, while the binding patterns and interaction mechanisms between novel protein aggregates such as amyloid fibrils (SAFs) formed through structural rearrangement and aroma compounds (especially homologues with the same parent nucleus) remain unclear. Therefore, this embodiment focuses on pyrazine homologues with increasing methyl substitution levels as aroma molecule probes to systematically reveal the structure-activity relationship between SAFs and aroma molecules, aiming to elucidate the influence mechanism of methylation degree on protein-aroma binding affinity.

[0106] The study found that all selected pyrazine aroma compounds could bind efficiently to SAFs-18. Figure 9 (A) Among them, SAFs-18 showed the highest binding rate to 2,3,5-trimethylpyrazine (38.40%), followed by 2,5-dimethylpyrazine (30.96%) and 2-methylpyrazine (21.34%). This result clearly shows that the binding of SAFs to pyrazine aroma compounds is significantly affected by the number of methyl substituents on the pyrazine ring, and the more methyl groups, the stronger the binding effect.

[0107] This phenomenon may stem from the synergistic changes in various non-covalent interactions between different methylated pyrazine aroma compounds and SAFs, leading to several hypotheses regarding different intermolecular forces. First, as the number of methyl groups increases, the overall hydrophobicity of the aroma molecule improves, potentially making it easier to embed into the binding regions composed of hydrophobic amino acid residues on the SAF surface, thereby enhancing hydrophobic interactions. Second, larger polymethylpyrazine aroma compounds may form broader contacts with the protein interface, resulting in stronger van der Waals attraction. Furthermore, polar groups exposed on the protein surface (such as amino and carboxyl groups) may interact with the nitrogen atoms in the pyrazine ring through hydrogen bonds, thereby improving the specificity and stability of the binding. However, the specific contribution weights of these different forces and their dominant mechanisms remain unclear and require further investigation.

[0108] 3.5.2. Aroma binding thermal stability The thermal stability of the aroma binding ability of SAFs-18 was evaluated by studying the binding behavior of SAFs-18 with three pyrazine aroma compounds at different temperatures. The results showed that the binding rate of pyrazine aroma compounds with different numbers of methyl groups to SAFs-18 decreased significantly with increasing temperature. Figure 9 BC in the middle; p <0.05 indicates that the binding ability is negatively correlated with temperature. Furthermore, regardless of the test temperature, the aroma binding ability remains positively correlated with the number of methyl substituents on the pyrazine ring. p <0.05).

[0109] From a thermodynamic perspective, the binding rate decreases with increasing temperature, suggesting that the binding process is exothermic (ΔH < 0). According to the principle of thermodynamic equilibrium, increasing the temperature will shift the chemical equilibrium towards the reverse reaction. In this system, the forward reaction is the binding of pyrazine aroma compounds to SAFs-18, and the reverse reaction is the dissociation of the bound pyrazine aroma compounds from SAFs-18. Therefore, increasing the temperature may promote dissociation and inhibit binding, thus leading to a decrease in the macroscopic binding rate. From a kinetic perspective, increasing the temperature increases the average kinetic energy of the molecules, making it easier for pyrazine aroma compound molecules bound to the hydrophobic cavity or surface of SAFs-18 to overcome the energy barrier and dissociate, i.e., increasing the dissociation rate constant, resulting in a decrease in the apparent binding rate. From a protein structure perspective, high temperature may induce structural "annealing" of amyloid fibers, which can lead to a more compact stacking of the amyloid fiber core; this rearrangement of the microstructure may affect the structure and exposure of hydrophobic regions in the protein aggregates, reducing their compatibility with pyrazine aroma compounds.

[0110] Example 3: A method for binding pyrazine aroma compounds with different methyl positions using pea protein amyloid fibers. 1. Preparation of PAFs with different acid heat treatment times A certain amount of PPI powder was added to 10 times its volume of deionized water and magnetically stirred at 25°C for 12 hours to ensure homogeneity. The pH of the solution was adjusted to 2.0 with 6 mol / L hydrochloric acid, and the mixture was magnetically stirred for 30 minutes. The solution was centrifuged (10000g, 4°C) for 30 minutes, and the supernatant was heated in an 85°C water bath for 6 hours, 12 hours, and 18 hours, respectively. After heating, the solution was immediately cooled to room temperature and stored at 4°C for at least 12 hours to prepare pea protein amyloid fibers (PAFs) with different hydrolysis times, named PAFs-6, PAFs-12, and PAFs-18.

[0111] 2. Test Characterization 2.1 Characterization of PAFs under different acid heat treatment times 2.1.1 The microstructure is the same as 2.1.1 in Example 1.

[0112] 2.1.2 The structural features are the same as those in Example 1, section 2.1.2.

[0113] 2.1.3 The physicochemical properties are the same as those in 2.1.3 of Example 1.

[0114] 2.1.4 Yield of PAFs under different acid heat treatment times PAFs dispersions were prepared according to the experimental steps in section 1, with water bath heating times set to 6 h, 12 h, 18 h, and 24 h, respectively. After acid heat treatment, cooling and hydration were performed to form mature PAFs dispersions, which were then freeze-dried to obtain PAFs powder, which was accurately weighed. The yield of PAFs was defined as the percentage of the mass of freeze-dried PAFs powder to the mass of the initially added PPI powder, calculated according to formula 3.1: Formula 3.1; Where: m PAFs Mass (g) of PAFs powder obtained after freeze-drying; m PPI : Initial mass of PPI powder (g).

[0115] 2.2 Preparation of PAFs-pyrazine aroma embedding complex PAFs were dispersed in phosphate buffer (10 mmol / L, pH 7.2), and three aroma compounds were dissolved in methanol (chromatographic grade) to prepare aroma substance stock solutions. Appropriate amounts of PAF solutions were mixed thoroughly with the different aroma substance stock solutions to achieve a final concentration of 0.1 w / v% for peptidoglycan and 0.08 mmol / L for aroma substances. A control group without added PAFs was included. The mixed solutions were incubated at 37°C in the dark for 16 h to allow for complete binding of PAFs with the aroma substances.

[0116] 2.2.1 The determination of aroma binding rate is the same as in 2.2.1 of Example 1.

[0117] 2.2.2 The evaluation of aroma binding thermal stability is the same as in 2.2.2 of Example 1.

[0118] 2.2.3 Analysis of the molecular interactions between PAFs and pyrazine aromas (1) Ultraviolet spectrum PAFs were dispersed in phosphate buffer to achieve a final concentration of 0.1% w / v for protein amyloid fibrils. Three aroma compounds were added, and the UV spectrum of the complex was scanned (200-400 nm) on a UV spectrophotometer (UV-2600i, Shimadzu, Japan).

[0119] (2) Synchronous fluorescence PAFs were dispersed in phosphate buffer to a final concentration of 0.1 w / v%. Three aroma compounds were added to final concentrations of 0, 0.10, 0.15, and 0.20 mmol / L. Fluorescence spectra were measured using a fluorescence spectrophotometer (FL6500, PerkinElmer, USA). The slit width was 10 nm, and the emission and excitation wavelengths were both 290 nm.

[0120] (3) Three-dimensional fluorescence The three-dimensional fluorescence intensity of the aroma-embedded complex was scanned using a fluorescence spectrophotometer (FL 6500, PerkinElmer, USA). Test parameters: excitation wavelength 190 nm, 32 scans, excitation wavelength interval 5 nm. Initial emission wavelength was 200 nm, final emission wavelength was 500 nm. Both the excitation and emission slits were 5 nm.

[0121] (4) Fluorescence thermodynamics Aroma-embedded complexes were incubated at different temperatures (298K, 308K, and 318K) for 30 min, and the fluorescence spectra of the samples were measured using a fluorescence spectrometer. The slit width for both excitation and emission was 5 nm, the excitation wavelength was 280 nm, and the emission spectral scanning wavelength was 300-400 nm. The mechanism of aroma quenching PAFs was determined by Stern-Volmer equation 3.2: Formula 3.2; In the formula, F0 and F are the fluorescence intensities of the blank control and the sample, respectively, [Q] is the concentration of aroma substances, and K is the fluorescence intensity of the sample. q τ is the quenching rate constant, and τ0 is the mean half-life of fluorescence (typically 10). -8 s), K SV This is the Stern-Volmer dynamic quenching constant.

[0122] For static quenching, Equation 3.2 is transformed into Equation 3.3 to obtain the binding constant (Ka) and the number of binding sites (n): Formula 3.3; The enthalpy change was further determined using the van't Hoff equation. H), entropy change ( S), change in free energy ( G) and other thermodynamic parameters, see Equations 3.4 and 3.5 for details: Formula 3.4; G= HT S Formula 3.5; Where R is the gas constant (8.314 J / K / mol) and T is the temperature (298 K, 308 K and 318 K).

[0123] (5) Molecular docking Molecular docking was performed using AutoDock software. Basic units of three aroma compounds were downloaded from the PubChem database (https: / / pubchem.ncbi.nlm.nih.gov) as ligands, and PAFs (PDB ID: 3SGO) were selected as acceptors. Protein structures were downloaded from a protein database (http: / / www.rcsb.org). During docking, the Lamarck genetic algorithm was used to perform 20 docking experiments on each of the three aroma compounds. The docking simulation result with the lowest docking energy was selected to represent the most favorable binding mode predicted by the program, thus clarifying the most likely binding mode between PAFs and aroma compounds. The optimal conformation was then visualized and analyzed using Pymol and Discovery Studio 4.5 software.

[0124] 2.2.4 Data Statistics and Analysis Methods Pearson correlation analysis was used to model the correlation between indicators, and correlation plots were generated using Origin 2024 software. Experiments were repeated three or more times, and results are expressed as mean ± standard deviation. One-way ANOVA and Tukey's test were performed using Statisticx 8.0 software. p Significance analysis was performed using <0.05, and graphs were generated using Origin 2024. Different letters in the graphs and tables represent statistically significant differences between groups. p <0.05).

[0125] 3. Results and Analysis 3.1. Microstructure of PAFs Natural pea protein has a globular conformation, but heating under acidic conditions (pH 2) induces time-dependent fibrosis. The degree of fibrosis in pea protein gradually increases with prolonged acid-heat treatment time (6–18 h). Figure 10 AC in the context of length distribution ( Figure 11 The average length of the resulting PAFs gradually increased from 111.107 nm (6 h) to 135.467 nm (12 h) and 211.274 nm (18 h), indicating that prolonged acid heating provided more time for longitudinal aggregation and growth of protein molecules, promoting continuous fiber elongation. Diameter distribution showed that the diameters of PAFs-6 (10.271 nm), PAFs-12 (8.330 nm), and PAFs-18 (6.267 nm) gradually narrowed with increasing heating time. Figure 11The results indicate that prolonged acid heating inhibited excessive lateral aggregation of protein molecules. This may be because intermolecular hydrogen bonds and hydrophobic interactions tend to be more ordered under acid heating conditions, driving the fibers towards a narrower diameter. The aspect ratios of PAFs-6 (11.855), PAFs-12 (18.592), and PAFs-18 (36.943) also showed an increasing trend. Figure 11 This indicates that extending the acid-heating time helps to construct "long and thin" protein amyloid fibers with a high aspect ratio. This structural evolution is closely related to the dynamic assembly of the protein fibrillation process: in the early stage of heating, protein molecules initially aggregate to form short fibers; as the acid-heating time increases, the fibers extend longitudinally, while lateral aggregation is restricted by ordered interactions, eventually forming fibrous assemblies with a high aspect ratio.

[0126] 3.2. Structural characteristics of PAFs 3.2.1. Cross-β-fold structure Thiofuric T staining is the "gold standard" for identifying amyloid fibrils. Its principle lies in the selective binding of thiosulfate T to the cross-β-sheet structure of amyloid fibrils, causing conformational changes in the dye and internal charge transfer, resulting in enhanced fluorescence. Compared to PPI, stained PAFs exhibit higher fluorescence intensity at 490 nm; and the fluorescence intensity gradually increases with prolonged acid heat treatment time (e.g., D in 10). This may be because prolonged acid heat treatment time exacerbates the formation of cross-β-sheet structures. This trend may be related to the fact that prolonged acid heat treatment promotes the formation of cross-β-sheet structures: after thiosulfate T binds to these structures, the stability of the molecule in the excited state is improved, and the fluorescence quantum yield increases accordingly, resulting in a stronger fluorescence signal. Therefore, the enhanced fluorescence of thiosulfate T provides direct evidence that prolonged acid heat treatment time leads to an increase in the content of cross-β-sheet structures after fibrillation, further confirming that pea protein undergoes a time-dependent fibrillation process under acidic conditions.

[0127] 3.2.2. Crystal Structure The protein sample exhibited distinct amorphous broad diffraction peaks at 8.72° (corresponding to an interplanar spacing of approximately 10.14 Å) and 20.04° (corresponding to an interplanar spacing of approximately 4.43 Å). Figure 10The broad peak near 8.72° typically corresponds to the α-helical conformation, while the broad peak near 20.04° reflects the characteristic signal of the amorphous or low-order structure of the protein, indicating that the packing and arrangement of molecular chains has limited regularity. From the PPI to the PAF series samples, the positions of these two amorphous broad diffraction peaks did not shift significantly, indicating that the basic amorphous structural framework of the protein molecule was not completely destroyed during the acid-heat-induced fibrillation process. However, the intensity of the amorphous broad diffraction peaks in PAFs decreased slightly compared to PPI, indicating that the original amorphous and α-helical conformations of the protein molecule underwent a certain degree of deconstruction, and some molecular chains may have transformed from loose amorphous or helical conformations to a more ordered fibrillated structure. Notably, PAFs showed new sharp diffraction peaks at 31.72° and 45.52°, and the intensity gradually increased with the extension of acid-heat treatment time, proving that acid-heat-induced fibrillation modification can promote the increase of PAF crystallinity and the ordering of the protein fibrillated structure.

[0128] 3.2.3. Protein Spatial Conformation Protein secondary structure: In FTIR spectroscopy, the absorption peak in the amide A region extends from 3000-3500 cm⁻¹. -1 The peak area and intensity of PAFs in the amide A region gradually increase with increasing acid hydrolysis time. This is caused by the stretching vibrations of NH and OH groups at the ions, and their intensity is closely related to hydrogen bonding interactions. Figure 10 The F band (1600-1700 cm⁻¹) indicates that prolonged acid heat treatment may enhance intermolecular hydrogen bonding interactions, which is beneficial to the stability and ordered arrangement of protein conformation. -1 The PAF (Protein Fiber Optic) region is mainly related to the stretching vibrations of CO and CN, and is the primary region for analyzing protein secondary structure. Compared to PPI, PAFs are located at 1600-1640 cm⁻¹. -1 The absorption peak intensity within the range increased significantly with prolonged acid heat treatment time. This result indicates that acid heat treatment induces a significant reconstruction of the secondary structure of PPI, driving the transformation of PPI from its natural amorphous or partially ordered conformation to a highly ordered amyloid fibrous structure.

[0129] Further quantitative analysis yielded results, as shown in Table 5.

[0130] Table 5 Secondary structure content of PPIs and PAFs

[0131] Table 5 shows that β-folds (1600-1640 cm⁻¹) -1The relative content of PPI gradually increased from 33.06% to 34.56% (PAFs-6), 36.10% (PAFs-12), and 36.22% (PAFs-18). This is likely because acid-heat conditions disrupted the native conformation of PPI, prompting peptide chains to refold and stack oriented via hydrogen bonds to form β-sheets, and prolonged acid-heat treatment further enhanced this ordered assembly process. Correspondingly, the relative proportions of random coils and β-turns gradually decreased. This may be because hydrolysis disrupted the local rigid conformation of the native protein, prompting some turns to dissociate and recombine into a more stable β-sheet structure, indicating that prolonged acid-heat treatment promoted the transformation of these two flexible structures in the protein molecule into β-sheet structures. In contrast, α-helices (1650-1660 cm⁻¹) showed a significantly higher proportion of random coils. -1 ) and intermolecular β-sheets (1620-1630 cm) -1 The relative content of ) did not change significantly. p The value >0.05 indicates that the significant increase in β-sheet content is not due to the unwinding of α-helices or intermolecular β-sheet transitions, but is mainly driven by the structural rearrangement of random coils and β-turns. This structural evolution pattern is consistent with the classic mechanism of protein amyloid fibrosis: the flexible regions of natural proteins (random coils, β-turns) refold under induced conditions into highly ordered β-sheet sheets, eventually assembling into amyloid fibers.

[0132] Synchronous fluorescence spectroscopy: Synchronous fluorescence spectroscopy can selectively detect changes in the microenvironment of tyrosine (Δλ=15nm) and tryptophan (Δλ=60nm) residues by using a fixed wavelength difference (Δλ). Figure 10 (GH in the text). With prolonged acid heat treatment, the characteristic peak of tyrosine gradually blue-shifted from 289.1 nm (PPI) to 284.2 nm (PAFs-18), indicating a decrease in the polarity of its surrounding microenvironment, suggesting that tyrosine residues may have migrated from the hydrophilic region to the hydrophobic environment. Correspondingly, the characteristic peak of tryptophan gradually red-shifted from 281.8 nm (PPI) to 282.7 nm (PAFs-18), indicating a slight increase in the polarity around tryptophan residues, possibly due to increased exposure to the hydrophilic environment. Furthermore, the fluorescence intensity of both decreased significantly, which may be due to the spatial confinement or energy transfer effect of the ordered assembly of the amyloid fibrous structure on the fluorescent group. These changes are consistent with the conformational rearrangement of protein amyloid fibrosis: the flexible conformation of the native protein gradually transforms into a highly ordered fibrous structure, prompting the migration of the microenvironment of aromatic amino acid residues from the polar hydrophilic environment to the hydrophobic fibrous interior.

[0133] Second-order ultraviolet (UV) spectroscopy: UV second-order spectroscopy can sensitively characterize tertiary conformational changes of proteins by observing the characteristic absorption peak shifts of tyrosine and tryptophan residues, as well as the ratio of the positive to negative absorption peaks (r=a / b). The four groups of samples exhibited positive absorption peaks near 288 nm and 295 nm, and negative absorption peaks near 283 nm and 291 nm. Figure 10 (I) The UV positive absorption peak at 288 nm is due to the combined effects of tyrosine and tryptophan, while the positive absorption peak at 295 nm represents only a change in tryptophan residues. Both of these UV positive absorption peaks showed a slight blue shift after prolonged hydrolysis time, indicating that fibrillation promoted the exposure of tryptophan residues from a hydrophobic environment to a hydrophilic environment. Simultaneously, the r-value, reflecting the relative microenvironmental polarity of tyrosine and tryptophan, gradually increased from 1.31 to 1.98, indicating a significant increase in solvent polarity around tyrosine residues, leading to the migration of more tyrosine residues to the hydrophobic region. In summary, with prolonged acid-heat treatment, the tertiary structure of the protein gradually unfolds, tyrosine residues shift to the hydrophobic environment, and tryptophan residues become more exposed to the hydrophilic environment, ultimately leading to an increase in system polarity and a corresponding rise in the r-value.

[0134] 3.2.4. Thiol group Free thiol groups refer to free thiol groups in proteins that do not participate in disulfide bond formation, while total thiol groups include both free thiol groups and those that participate in disulfide bond formation. Both reflect the exposure and oxidation state of thiol groups in proteins, playing a crucial regulatory role in protein conformational stability, aggregation tendency, and functional properties. With prolonged acid heat treatment, the total thiol content gradually increased from 0.666 μmol / g (PPI) to 0.721 μmol / g (PAFs-6), 0.939 μmol / g (PAFs-12), and 1.345 μmol / g (PAFs-18). Figure 12 (A) This may be because acid heat treatment induces protein molecule unfolding, gradually exposing the previously encapsulated thiol groups to the molecular surface. The free thiol content gradually increased from 0.379 μmol / g (PPI) to 0.519 μmol / g (PAFs-6), 0.796 μmol / g (PAFs-12), and 1.157 μmol / g (PAFs-18), possibly related to the breakage of some disulfide bonds during acid heat treatment. Notably, the difference between total thiol groups and free thiol groups (reflecting disulfide bond content) showed a trend of first decreasing and then stabilizing with treatment time. This may be because disulfide bond breakage is dominant in the early stages of treatment, leading to an increase in free thiol groups; as the acid heat treatment time prolongs, the exposed thiol groups and disulfide bond rearrangement gradually reach a dynamic equilibrium, causing the difference to stabilize.

[0135] 3.2.5. Hydrophobic groups With the extension of acid heat treatment time, the surface hydrophobicity of PAFs shows an increasing trend.p <0.05) Figure 12 (B in the text). This is mainly because acid-heat conditions disrupt the native conformation of proteins, causing their originally tight globular structure to gradually denature and unfold, and the peptide chains to unfold. This exposes the hydrophobic groups that were originally encapsulated within the molecule, providing numerous binding sites for the 8-aniline-1-naphthalenesulfonic acid probe, ultimately resulting in a significant increase in fluorescence intensity. Furthermore, longer acid-heat treatment provides sufficient time for conformational rearrangement to allow for continued exposure of hydrophobic groups, further enhancing the hydrophobic properties of PAFs. This trend of gradually increasing surface hydrophobicity is consistent with the mechanism of hydrophobic interactions driving the orderly stacking of β-sheets during protein amyloid fibrosis, and also provides a structural basis for the orderly assembly of PAFs and the formation of their functional properties.

[0136] 3.2.6. SDS-PAGE The electrophoretic pattern of PPI shows clear protein subunit bands. Figure 12 C in the text corresponds to 7Sα. ， The PAFs contained 7Sα (~75kDa), 7Sβ (~63kDa), and 7Sβ (~48kDa) subunits, as well as 11S acidic (29-33kDa) and basic (18-22kDa) subunits. Subunit bands greater than 20kDa in PAFs gradually weakened or even disappeared with acid heating time, while a large number of low molecular weight aggregates appeared in the 5-20kDa range. Combined with optical density analysis (Table 6), it was found that the high molecular weight peptide (29-98kDa) bands in PPIs completely disappeared after fibrinization treatment, while the optical density of the medium molecular weight polypeptide (18-22kDa) bands significantly increased. p <0.05), and new bands appeared in the low molecular weight region (5-11 kDa). These results indicate that acid-thermal treatment induces the dissociation and degradation of the 7S and 11S subunits in PPI, generating small polypeptides. Subsequently, these polypeptides, driven by hydrogen bonding and hydrophobic interactions, directionally stack and gradually assemble into β-sheet-rich amyloid fibers. The progressive hydrolysis and orderly aggregation of protein subunits in this process are key steps in amyloid fiber formation, providing direct molecular-level evidence for the structure and functional properties of PAFs.

[0137] Table 6. Absolute quantitative ratios of PPI and PAFs solution electrophoresis bands

[0138] Note: "—" indicates that no electrophoretic bands were detected. The data is the average of three independent replicates. Different letters (ad) in the same row indicate that the difference between groups is statistically significant. p <0.05).

[0139] 3.3. Physicochemical properties of PAFs 3.3.1. Potential The zeta potential reflects the net charge characteristics of the protein particle surface. The zeta potential of PPI is -35.46 mV, and it is generally negatively charged. Figure 13 (A) After acid-heat treatment, the Zeta potential of PAFs turned positive and gradually increased with the extension of treatment time. This reversal of potential from negative to positive may be due to the pH of the solution containing the protein being much lower than the isoelectric point of pea protein isolate, at which point the dissociation of carboxyl groups is inhibited, while the amino groups are fully protonated. Notably, the Zeta potential of PAFs continued to increase with the extension of acid-heat treatment time, which may be related to the further unfolding of the peptide chain and the exposure of more positively charged groups. The above results indicate that acid-heat treatment helps to improve the colloidal dispersion stability of proteins by regulating the surface charge characteristics, providing an interfacial chemical basis for the application of PAFs in food colloidal systems and functional delivery.

[0140] 3.3.2. Turbidity and Particle Size Turbidity and particle size are key indicators characterizing protein aggregation states. PPI has the highest turbidity and particle size. Figure 13 (B in the text), while the turbidity and particle size of PAFs-6 decreased by 42.3% and 57.2% respectively compared to PPI (in the text). p <0.05). This change is mainly due to two factors: 1) the H+ provided by hydrochloric acid. + 1) The natural spatial structure of PPIs was disrupted, promoting peptide chain breakage and exposing hydrophobic regions; 2) The acidic environment (pH 2.0) caused the net charge on the protein surface to change from negative to positive (Zeta potential increased to +29.77mV), which was much higher than its isoelectric point. The enhanced electrostatic repulsion effectively inhibited particle aggregation, resulting in a reduction in particle size. Notably, the particle size of PAFs-12 and PAFs-18 showed a reversal trend compared to PAFs-6. p <0.05). This may be because the acid-heat environment further induces conformational rearrangement of the peptide chains, and the exposed hydrophobic regions drive peptide chain self-assembly through hydrophobic interactions, forming fibrous aggregates with a cross-beta-sheet structure. At the same time, the extended acid-heat time also promotes the continuous growth and fusion of fibers, ultimately leading to an increase in aggregate size. In addition, the turbidity of protein solutions is usually positively correlated with aggregate size. Therefore, the significant increase in particle size of PAFs after acid-heat treatment can be explained as an increase in turbidity caused by the increase in aggregate size in the solution during the early stage of fibrinization.

[0141] 3.3.3. Dispersion Stability Polydispersity index (PDI) is a key parameter characterizing the uniformity of particle size distribution in a particulate system. The PPI value of PAFs decreases significantly with increasing acid heat treatment time. p <0.05; Figure 13(D in the text). This is likely because the acid-thermal environment induces the unfolding of protein peptide chains, exposing hydrophobic regions, which then self-assemble into uniform fibrous aggregates under the synergistic drive of hydrophobic interactions and β-sheet hydrogen bonds. Simultaneously, the zeta potential on the protein surface continuously increases (from negative to positive and gradually rises), and the enhanced electrostatic repulsion effectively inhibits the random aggregation of particles, thus making the particle size distribution of the PAF system more concentrated. This change not only reflects the transformation process of PAFs from disordered aggregation to ordered fibrous structure driven by acid-thermal treatment, but also confirms from the perspective of particle size uniformity that the stability of the PAF dispersion system is significantly improved with fibrous modification.

[0142] 3.3.4. Solubility Protein solubility is the outward manifestation of the thermodynamic equilibrium reached between protein-protein interactions and protein-solvent interactions. Its variation is determined by both average hydrophobicity and net surface charge: hydrophobic interactions promote protein-protein aggregation, reducing solubility; ion hydration enhances the affinity between protein and water, thus increasing solubility. The solubility of PPI is 38.2 mg / mL, while the solubilities of PAFs-6, PAFs-12, and PAFs-18 decrease to 28.5 mg / mL, 19.8 mg / mL, and 13.5 mg / mL, respectively. Figure 13 C in the text). Although the absolute value of the Zeta potential (net surface charge characteristic) increases after acid heat treatment ( Figure 13 In the formula (A), the solubility can theoretically be improved by enhancing ion hydration; however, the surface hydrophobicity increases significantly with treatment time (PPI is 2.75 × 10⁻⁶). 6 CPS, PAFs-18 increased to 9.04 × 10 6 The CPS (Chemical Particle Size Analysis) indicates that acid heat induces the exposure of a large number of hydrophobic groups, which strengthens the hydrophobic interactions between proteins. Under these conditions, the inhibition of solubility by hydrophobic interactions dominates, ultimately leading to a significant decrease in the solubility of PAFs with prolonged treatment time.

[0143] 3.3.5. Thermal stability The conformational thermal stability of PPIs and PAFs can be analyzed using thermogravimetric curves and thermogravimetric differential curves. Their thermal degradation process mainly consists of two stages. The initial stage (<150℃) of weight loss primarily originates from water evaporation. During this stage, the highest thermal degradation temperature for PPI is 40.66℃, while for PAFs-6, PAFs-12, and PAFs-18 it increases to 51.73℃, 56.56℃, and 59.46℃ respectively. Figure 13 (E in the text). This indicates that the binding capacity of protein molecules to water molecules increases with prolonged acid heat treatment time, which may be related to the gradual adjustment of protein conformation to form more stable hydration sites.

[0144] The mass loss in the second stage (>150℃) is mainly attributed to the thermal degradation of the protein's structural backbone. In this stage, the maximum thermal degradation temperature increases sequentially from 307.54℃ (PPI) to 323.38℃ (PAFs-6), 330.97℃ (PAFs-12), and 332.91℃ (PAFs-18); simultaneously, the mass loss rate decreases sequentially from 68.60% (PPI) to 61.65% (PAFs-6), 58.50% (PAFs-12), and 56.19% (PAFs-18), indicating that extending the acid heat treatment time can improve the conformational thermal stability of native PPI. This is because prolonged heating increases the proportion of ordered structures such as β-sheets in the secondary structure of the protein, and β-sheet structures can form a highly ordered stacked network through intermolecular hydrogen bonds; at the same time, the hydrophobic groups exposed by acid heat treatment further strengthen intermolecular bonds through hydrophobic interactions, resulting in PAFs forming a more compact and more force-rich aggregated structure. This change aligns with the formation mechanism of amyloid fibers: during fiber assembly, extensive hydrogen bonds and hydrophobic interactions between peptide side chains construct a stable supramolecular network, thereby enhancing the polymer's thermal stability, manifested as an increase in maximum thermal degradation temperature and a decrease in mass loss rate.

[0145] 3.3.6. Rheological Behavior The rheological behavior of proteins is closely related to their gel quality, with storage modulus (G', representing solid behavior) and loss modulus (G", representing liquid behavior) being important indicators of gel viscoelasticity. G' > G" generally indicates that the system exhibits solid-state-dominated gel properties. At the same angular frequency, the G' values ​​of all four samples were greater than the G" values ​​( Figure 13 The FG and G′ values ​​of the PAFs gel both increased with increasing frequency (0.1~100Hz), indicating the formation of a continuous and stable three-dimensional network structure. With prolonged acid heat treatment, the G′ and G′′ values ​​of the PAFs gel significantly increased and were both higher than the PPI. This phenomenon can be attributed to the full fibrillation and conformational rearrangement of the protein molecules: acid heat treatment improved the orderliness of the secondary structure, increased intermolecular cross-linking points, and strengthened network entanglement, thereby constructing a denser and more robust three-dimensional gel network.

[0146] Furthermore, all four protein solutions exhibited shear-thinning behavior, meaning that the apparent viscosity decreased with increasing shear rate. Figure 13The H in the text conforms to the characteristics of a non-Newtonian fluid. At low shear rates, the entanglement and interactions between molecular chains are well maintained, resulting in high flow resistance and viscosity. As the shear rate increases, external forces orient protein molecules along the shear direction and disentangle them, disrupting some intermolecular interactions, reducing fluid resistance, and consequently decreasing viscosity. Among these, PPI exhibits the lowest overall viscosity and a relatively gentle shear-thinning trend; while PAFs, due to prolonged acid heat treatment leading to more complete molecular cross-linking, form stronger network resistance (higher initial viscosity) at low shear rates. At high shear rates, the network structure is more significantly disrupted, enhancing molecular orientation and disentanglement effects, thus exhibiting a more dramatic decrease in viscosity.

[0147] 3.3.7. Scanning Electron Microscopy The PPI sample exhibits a relatively regular, continuous sheet-like structure with a relatively tight molecular arrangement. Figure 13 The presence of I in PAFs (part of the protein matrix) is related to the fact that they were not sufficiently heat-treated, resulting in relatively simple intermolecular interactions. However, the microstructure of the PAFs series samples obtained after acid heat treatment showed significant changes with heating time: the sheet-like structure of PAFs-6 began to break and become irregular, forming more pores and scattered fragments, indicating that even a short heating time had already induced preliminary conformational changes and aggregation remodeling of the protein molecules. PAFs-12 began to form small sheet-like structures with more complex interlayer connections and a wider pore distribution. This may be because with prolonged acid heat treatment, the protein molecules had more time for conformational adjustments and interactions.

[38] This process drives the gradual fusion and growth of small-scale sheets. The sheet structure of PAFs-18 further increases in size and thickness, forming more continuous and larger-scale sheet-like aggregates. This structural evolution is consistent with the molecular behavior of proteins under acid-heat conditions: heating time, as a key regulatory factor, promotes the rearrangement of protein secondary structures, driving molecules to continuously cross-link and aggregate through non-covalent interactions.

[0148] 3.4. Yield of PAFs Acid heat treatment time significantly affected the formation efficiency of pea protein amyloid fibers (PAFs). With prolonged heating time, the yield of PAFs showed a significant increasing trend. p <0.05). Specifically, the yield increased slowly from 30.35% at 6h to 35.35% at 12h; subsequently, it experienced a rapid growth phase between 12h and 18h, finally reaching 51.54% at 18h. Figure 13 (J in the text). This nonlinear growth conforms to the typical "nucleation-elongation" mechanism of amyloid fibril formation: in the initial stage of heating, acid heat induces the protein to unfold and expose hydrophobic residues, forming a fibril nucleus; then, in the rapid elongation phase, a large number of free protein monomers or oligomers rapidly assemble onto the fibril nucleus through hydrophobic interactions and hydrogen bonds, resulting in a significant increase in yield.

[0149] However, when the heating time was further extended to 24 hours, the yield only increased slightly to 52.12%, which was not significantly different from that of 18 hours. p >0.05). This indicates that by 18 h, the available protein monomers in the system had been almost completely consumed, and the fibrinization reaction had reached kinetic equilibrium or saturation. Excessive heating (24 h) did not significantly promote further fiber accumulation; instead, it may have caused some of the formed fibers to depolymerize or hydrolyze. Considering both preparation efficiency and yield, as well as the structural damage that excessive heating may cause, this example ultimately selected 18 h as the optimal heating time for preparing PAFs for subsequent experiments.

[0150] 3.5. Binding ability of PAFs to pyrazine aromas with different methyl positions 3.5.1. Aroma Binding Rate In recent years, the influencing factors of protein-aroma binding have received widespread attention. Compared with the type and structure of proteins, the structural characteristics of aroma molecules themselves may be more dominant in regulating binding behavior. Based on the speculation that the methyl position may have a key influence on protein-aroma binding, this example selects three pyrazine compounds with different methyl positions: 2,5-dimethylpyrazine (para), 2,6-dimethylpyrazine (meta), and 2,3-dimethylpyrazine (ortho), and systematically investigates their binding behavior with PAFs. At 37°C, the binding rates of PAFs-18 with 2,5-dimethylpyrazine, 2,6-dimethylpyrazine, and 2,3-dimethylpyrazine were 31.1%, 28.5%, and 24.7%, respectively. p <0.05; Figure 14 The A group exhibits a clear "para > meta > ortho" pattern. This result confirms that the methyl substitution position on the pyrazine ring is a key structural factor regulating the binding behavior of pyrazine aroma compounds to protein amyloid fibers.

[0151] This phenomenon can be explained by the synergistic regulation of multiple intermolecular forces. 1) Steric hindrance is the core dominant factor: Compared to the ortho and meta isomers, the methyl position of 2,5-dimethylpyrazine (para position) minimizes its steric hindrance, making it easier to embed into the hydrophobic pocket of PAFs; while 2,3-dimethylpyrazine, due to the significant ortho steric hindrance, hinders its entry into the binding site, indicating that the spatial arrangement of the methyl group directly restricts its binding affinity to the protein. 2) Structure dependence of hydrophobic interactions: The methyl position alters the spatial distribution of the hydrophobic region of the pyrazine ring. 2,5-dimethylpyrazine has a better match with the hydrophobic site of PAFs-18, forming a stronger hydrophobic interaction; conversely, the steric hindrance of 2,3-dimethylpyrazine restricts the full contact between its hydrophobic region and the protein surface, resulting in a weakened hydrophobic interaction. 3) Synergistic stabilizing effect of van der Waals forces and hydrogen bonds: The relatively open conformation of 2,5-dimethylpyrazine provides a larger van der Waals contact area and better polarity matching, which is conducive to the formation of stable hydrogen bonds; while the spatially restricted conformation of 2,3-dimethylpyrazine weakens its contact area and bonding ability with the protein interface, thereby reducing the binding rate. In summary, the methyl position may systematically regulate the binding strength of pyrazine aroma molecules and PAFs by adjusting intermolecular forces such as steric hindrance, hydrophobic interactions, and hydrogen bonds, but the specific regulatory mechanism needs further confirmation.

[0152] 3.5.2. Aroma binding thermal stability Temperature significantly affected the binding ability of PAFs-18 to the three pyrazine isomers. As the ambient temperature increased from 37℃ to 120℃, the binding rates of all three aroma compounds showed a significant decreasing trend. p <0.05). Specifically, under simulated grilling or baking conditions at 120°C, the binding rates of PAFs-18 to 2,3-dimethylpyrazine, 2,6-dimethylpyrazine, and 2,5-dimethylpyrazine decreased to 8.21%, 13.37%, and 14.83%, respectively. Figure 14 (BC in the text). This decrease may be attributed to localized loosening of the protein structure caused by high temperature, impaired integrity of the hydrophobic pocket, reduced effective binding sites, and easier breakage of hydrogen bonds due to increased energy; at the same time, increased volatility of aroma substances may also affect binding performance, but weakened binding forces are still likely the dominant factor.

[41] .

[0153] It is noteworthy that although high temperatures significantly reduced the overall binding strength, the binding ability of pyrazine isomers remained consistent across temperature gradients: 2,5-dimethylpyrazine (para) > 2,6-dimethylpyrazine (meta) > 2,3-dimethylpyrazine (ortho). This consistent stability across temperature levels further confirms that steric hindrance may be the core structural factor determining the affinity of PAFs for aroma binding. This finding is consistent with previous research, where the spatial structure of aroma molecules and the position of substituents directly determine their ease of entry into the hydrophobic cavity of proteins; aroma molecules with less steric hindrance tend to exhibit higher affinity. In this embodiment, 2,5-dimethylpyrazine, with its minimal steric hindrance and superior linear conformation, maintained the most stable binding even under thermal perturbations caused by high temperatures. In contrast, the greater ortho-steric hindrance of 2,3-dimethylpyrazine made it difficult for it to penetrate deep into the protein binding site. This structural binding barrier could not be eliminated by temperature changes, resulting in its consistently lowest binding rate across all test temperatures.

[0154] 3.6. Molecular interactions between PAFs and pyrazine aromas with different methyl positions 3.6.1. Ultraviolet Spectrum Ultraviolet absorption spectroscopy reveals the interactions between proteins and small molecule ligands by measuring changes in the microenvironment in which complexes form. For example... Figure 14 As shown in Figure D, PAFs-18 exhibits a smooth UV absorption curve in the 240-400 nm range with no obvious characteristic peaks. The differences in UV absorption intensity among the three pure pyrazine aroma compounds are negligible; however, after binding with PAFs-18, all complexes show a significant color-enhancing effect. Specifically, compared to the pure aroma compounds, the PAFs / 2,5-Dimethylpyrazine complex shows the greatest increase in absorbance (ΔAbs=0.332), followed by PAFs / 2,6-Dimethylpyrazine (ΔAbs=0.321) and PAFs / 2,3-Dimethylpyrazine (ΔAbs=0.266). This significant increase in absorbance, without a significant shift in the maximum absorption wavelength, strongly confirms that the interaction between PAFs-18 and pyrazine molecules is not a simple physical mixture, but rather an effective binding to form a stable ground-state complex.

[0155] 2,5-Dimethylpyrazine, as the para-isomer, possesses minimal steric hindrance and a highly symmetrical linear conformation, enabling it to better bind to the fibrous network structure of PAFs. This significantly promotes the formation of tight and extensive π-π stacking between the pyrazine ring and the aromatic amino acid residues of the protein, resulting in the highest macroscopic increase in absorbance. In contrast, 2,3-Dimethylpyrazine, due to its larger ortho-steric hindrance, limits its effective contact area and the amount of protein it binds. This hinders effective orbital overlap between it and the protein chromophore, leading to poorer stability or a smaller number of bound complexes, thus exhibiting the weakest chromophore effect. The difference in absorbance increase among the different isomers further reveals the crucial regulatory role of the steric hindrance effect of the methyl group in the protein-aroma binding process.

[0156] 3.6.2. Synchronous fluorescence and three-dimensional fluorescence Fluorescence spectroscopy further explored the protein microenvironment response, utilizing endogenous fluorophores (tryptophan and tyrosine) as highly sensitive molecular probes to reveal the specific perturbations of protein-aroma binding on the internal hydrophobic regions and spatial conformation of proteins. Synchronous fluorescence analysis showed that the introduction of all three aroma ligands led to significant quenching of the endogenous fluorescence of PAFs, indicating the successful binding of aroma substances and the alteration of the microenvironmental polarity around tyrosine and tryptophan residues. Figure 15 Three-dimensional fluorescence further confirms this result. Figure 16 ): Two characteristic fluorescent centers of PAFs—Peak A (λ) characterizing the tertiary structure ex / λ em nm=280 / 340nm) and π→π with the polypeptide backbone Peak B(λ) related to the transition ex / λ em =230 / 340 nm), the fluorescence intensity of both peaks decreased significantly after binding with aroma compounds, and the contour density was noticeably sparse. Notably, both spectroscopic techniques exhibited a consistent isomer-dependent fluorescence quenching pattern: the residual fluorescence intensity of the complex system consistently followed the order 2,3-dimethylpyrazine (ortho) > 2,6-dimethylpyrazine (meta) > 2,5-dimethylpyrazine (para). Furthermore, the subtle deformation of the three-dimensional fluorescence characteristic peak edges suggests that the protein peptide chain may have undergone local unfolding or flexible rearrangement during binding to accommodate the insertion of the hydrophobic ligand.

[0157] This cross-dimensional spectroscopic consistency confirms that methyl position-mediated steric hindrance is the core driving force regulating protein-aroma binding mechanisms. 2,5-Dimethylpyrazine (para position), with its highly symmetrical linear conformation and minimal steric hindrance, exhibits extremely strong molecular permeability; it can overcome the physical barrier of the PAF fibrous network, deeply embedding itself in the hydrophobic core of the protein, binding to deeply embedded tryptophan / tyrosine residues, thus exhibiting the strongest quenching effect. Conversely, the significant steric hindrance effect of the adjacent methyl group in 2,3-dimethylpyrazine (ortho position) restricts its diffusion ability in the protein matrix, hindering its approach to the chromophore deep within the hydrophobic core, causing it to mainly remain on the protein surface or in the hydrophilic region, resulting in looser binding and greater distance, thus retaining the highest residual fluorescence intensity.

[0158] 3.6.3. Fluorescence Thermodynamics To elucidate the interaction types and thermodynamic driving forces between PAFs-18 and the three pyrazine isomers, this example analyzes fluorescence quenching data at three different temperatures (298, 308, and 318 K). The results are as follows: Figure 17 As shown in Table 7.

[0159] Table 7. Fluorothermodynamic parameters of 2,3-methylpyrazine, 2,5-dimethylpyrazine and 2,6-trimethylpyrazine bound to PAFs.

[0160] Table 7 and Figure 17 The results show that the quenching type was determined using the Stern-Volmer equation. The Stern-Volmer quenching constant (K0) for each system is also shown. SV All of them exhibit a significant negative temperature dependence, i.e., K SV The rate decreases with increasing temperature. This trend strongly confirms that the quenching process is mainly caused by static quenching resulting from the formation of stable ground-state complexes between aroma substances and proteins, rather than dynamic collisions. Furthermore, the bimolecular quenching rate constant (K0) decreases with increasing temperature. q The order of magnitude of 10 is reached. 10 L·mol -1 ·s -1 Approaching or slightly exceeding the maximum dynamic collision limit of various quenchers (2.0 × 10⁻⁶). 10 L·mol -1 ·s -1 This further supports the dominance of the static quenching mechanism.

[0161] Secondly, the combination constant (K) was obtained through the double logarithmic equation. aThe number of binding sites (n) was also considered. The n values ​​for all systems were close to 1 (0.6–1.2), indicating that the pyrazine molecule and PAFs primarily maintained a 1:1 stoichiometric binding mode. Notably, the binding affinity (K0) of the three isomers to the protein was significantly different. a The binding constants of the isomers consistently exhibit a strong order of strength: 2,5-dimethylpyrazine (para) > 2,6-dimethylpyrazine (meta) > 2,3-dimethylpyrazine (ortho). As temperature increases, the intensified molecular thermal motion leads to a decrease in the binding constants of all systems, but the relative binding ability differences between isomers remain unchanged. This stable pattern across temperature ranges strongly confirms that the steric hindrance effect is not a random thermodynamic phenomenon, but a core physical characteristic determined by the inherent spatial configuration of the molecule: the para isomer, with its minimal steric hindrance, consistently grants it the ability to overcome thermal disturbances, penetrate the hydrophobic core, and maintain the strongest binding, while the significant steric hindrance of the ortho isomer consistently limits the tightness of its binding.

[0162] Finally, the thermodynamic parameters were calculated using the van 't Hoff equation to determine the main driving force of the bonding process. The Gibbs free energy change for all systems was calculated. G) are all negative (-12.4 to -24.5 kJ·mol⁻¹). -1 This indicates that the binding of PAFs to pyrazine molecules is a spontaneous process. Simultaneously, both the enthalpy change (ΔH) and entropy change (ΔS) are negative. According to thermodynamic criteria, when ΔH < 0 and ΔS < 0, it indicates that the intermolecular forces are mainly dominated by hydrogen bonds and van der Waals forces. Therefore, the binding of PAFs to pyrazine aroma compounds is a spontaneous process primarily maintained by hydrogen bonds and van der Waals forces and driven by enthalpy.

[0163] 3.6.4. Molecular docking This study selected PAFs as acceptors and three pyrazine isomers as ligands to further explore the interaction mechanism between aroma compounds and PAFs through molecular docking. The 2D interaction map shows that the aroma compounds were successfully embedded in the cross-β-sheet structure of PAFs (a typical structure of protein amyloid filaments), theoretically demonstrating that PAFs can effectively encapsulate pyrazine aroma compounds with different methyl positions. Figure 18 ).

[0164] Specifically, 2,5-dimethylpyrazine, with its highly symmetrical linear configuration, can deeply fit the cross-β-sheet structure. The nitrogen atom of the pyrazine ring forms a stable conventional hydrogen bond with Ala C:5; simultaneously, the aroma molecule forms extensive Pi-alkyl (Pi-Alkyl) interactions with Leu C:6, Leu A:6, Ala C:4, and Ala B:4, with van der Waals forces present at Ala B:5. Notably, 2,5-dimethylpyrazine also forms a specific Pi-Sigma interaction with Leu B:6. Although its energy intensity is lower than that of hydrogen bonds, this directional non-covalent contact makes a crucial contribution to anchoring ligand orientation and enhancing the stability of the complex within the hydrophobic core.

[0165] In contrast, the binding of 2,6-dimethylpyrazine to PAFs mainly relies on hydrogen bonds formed with Ala C:4 and Ala D:5, as well as Pi-alkyl interactions with Ala D:4, Leu C:6, and Leu D:6. It is also surrounded by van der Waals forces provided by residues such as Ala C:5, but lacks the support of strong Pi-Sigma interactions, resulting in a weaker binding tightness than 2,5-dimethylpyrazine.

[0166] For 2,3-dimethylpyrazine, the significant steric hindrance effect of its ortho-methyl group hinders the penetration of the ligand into the deep hydrophobic microenvironment. Although it can form hydrogen bonds with Ala D:4 and Ala E:5, and exhibit Pi-alkyl interactions with Leu E:6 and Ala E:4, and van der Waals forces with Ala D:5, the total number of binding sites is significantly reduced compared to 2,5-dimethylpyrazine and 2,6-dimethylpyrazine. This loss of key binding sites directly leads to a decrease in binding affinity. Molecular docking results confirm at the atomic level that the methyl position modulates its matching degree with the hydrophobic side chains of PAFs by altering the spatial geometry of the molecule, thereby determining the strength of the binding affinity.

[0167] 3.7. Methyl position mediates the binding mechanism of PAFs-pyrazine aromas This embodiment integrates macroscopic binding kinetic data, thermodynamic parameters, and microscopic molecular simulation evidence to construct a PAFs-pyrazine aroma binding mechanism model based on methyl substitution position regulation. Figure 19 Although the three pyrazine isomers have the same chemical composition, their binding affinity to PAFs exhibits a significant structure-activity dependence: 2,5-dimethylpyrazine > 2,6-dimethylpyrazine > 2,3-dimethylpyrazine.

[0168] The core of this binding mechanism lies in the "lock-key" match between the spatial geometry of the isomer molecule and the hydrophobic cavity microenvironment of PAFs, primarily driven by the steric hindrance effect mediated by the methyl position. 2,5-Dimethylpyrazine exhibits the best binding rate (31.1%) and thermal stability, mainly attributed to its unique para-substitution structure. First, geometrically, the para-methyl group endows the molecule with a highly linear symmetric structure. This flat, streamlined structure effectively overcomes the physical barrier posed by the dense fibrous network on the PAF surface, allowing for easier embedding into the deep hydrophobic pockets of the "cross-β" folded structure. Second, from the perspective of molecular forces, in addition to conventional hydrophobic interactions (Pi-Alkyl / Alkyl) and hydrogen bonds, the pyrazine ring of 2,5-dimethylpyrazine acts as a π-electron donor, forming a highly directional Pi-Sigma interaction with the PAF side chains.

[0169] In contrast, 2,3-dimethylpyrazine exhibits the weakest binding affinity (24.7%), primarily due to significant steric hindrance. The ortho-substitution results in a significant increase in volume on one side of the molecule, disrupting its linearity. This conformational asymmetry significantly limits its penetration into high-affinity sites within PAFs. This structural barrier leads to two negative consequences: first, limited binding depth, as 2,3-dimethylpyrazine struggles to penetrate high-affinity sites within PAFs, remaining primarily on the fiber surface through weak, non-specific adsorption. Second, a distorted binding conformation; the steric hindrance forces the pyrazine ring to deviate from the optimal binding angle, preventing the formation of Pi-Sigma-like interactions, resulting in lower binding energy and, macroscopically, poor thermal stability, making it prone to desorption and escape during heating.

[0170] The methyl groups of 2,6-dimethylpyrazine are distributed in the meta position, resulting in better molecular symmetry than 2,3-dimethylpyrazine, but less linearity. This spatial configuration allows it to enter some hydrophobic cavities, but it is slightly inferior to 2,5-dimethylpyrazine in terms of binding depth and the strength of the non-covalent network formed, thus exhibiting moderate binding rate (28.5%) and thermal stability.

[0171] In summary, the binding behavior of PAFs to pyrazine aromas follows a strict structure-activity relationship. The substitution position of the methyl group acts as a key structural switch, influencing the stability of aroma compounds bound to proteins by adjusting the steric hindrance of the molecule.

[0172] This embodiment systematically reveals the potential of PAFs as aroma carriers and their binding mechanism with pyrazine aroma compounds at different methyl positions. Overall, acid-thermal induction successfully drove the conformational remodeling of pea protein, leading to its self-assembly into PAFs possessing a high aspect ratio morphology, typical cross-β-sheet characteristics, and excellent surface hydrophobicity. The methyl substitution positions of pyrazine isomers significantly modulate their binding affinity with PAFs, exhibiting a strict structure-activity dependence of 2,5-dimethylpyrazine (para, 31.1%) > 2,6-dimethylpyrazine (me, 28.5%) > 2,3-dimethylpyrazine (ortho, 24.7%); and this affinity gradient ordering exhibits significant thermal stability, remaining consistent across a wide temperature range of 50–120 °C. This binding process is essentially a spontaneously enthalpy-driven process maintained by hydrogen bonds, van der Waals forces, and hydrophobic interactions. 2,5-Dimethylpyrazine, with its low steric hindrance due to its para-linear configuration, forms the richest intermolecular interaction network with amyloid fibers, significantly increasing the number of effective binding sites and forming specific Pi-Sigma interactions. In summary, this embodiment elucidates the binding mechanism of PAFs with pyrazine aromas at different methyl positions at the molecular interaction level, providing a solid theoretical basis for developing efficient plant-based flavor delivery systems based on differences in aroma molecule configurations.

[0173] The above description is only a preferred embodiment of the present invention. It should be noted that for those skilled in the art, several improvements and modifications can be made without departing from the principle of the present invention, and these improvements and modifications should also be considered within the scope of protection of the present invention.

Claims

1. A protein amyloid fiber-pyrazine aroma compound composite material, comprising protein amyloid fibers and pyrazine aroma compounds; The protein amyloid fibers include one or more of soybean protein amyloid fibers, pea protein amyloid fibers, rice protein amyloid fibers, and wheat protein amyloid fibers; The pyrazine aroma compounds include one or more of 2,5-dimethylpyrazine, 2,6-dimethylpyrazine, 2,3-dimethylpyrazine, 2-methylpyrazine, and 2,3,5-trimethylpyrazine.

2. The protein amyloid fiber-pyrazine aroma compound according to claim 1, characterized in that, The ratio of protein amyloid fibers to pyrazine aroma substances is 0.5~1.5g:0.05~0.1mmol.

3. The protein amyloid fiber-pyrazine aroma compound according to claim 1, characterized in that, The method for preparing the protein amyloid fibers includes the following steps: The protein powder and water are first mixed, and the resulting first mixture is adjusted to be acidic. Hydration and fibrosis are then carried out sequentially to obtain the protein amyloid fibers.

4. The protein amyloid fiber-pyrazine aroma compound according to claim 3, characterized in that, The mass ratio of the protein powder to water is 1:9~11, and the first mixing time is 10~14h.

5. The protein amyloid fiber-pyrazine aroma compound according to claim 3, characterized in that, The acidic pH value is 1.5~2.5, the hydration time is 20~35 min, the fibrosis temperature is 80~90℃, and the time is 6~20 h.

6. The method for preparing the protein amyloid fiber-pyrazine aroma compound according to any one of claims 1 to 5, characterized in that, Includes the following steps: Amyloid fibers were dispersed in a buffer solution, and then pyrazine aroma compounds were added. The resulting mixture was then equilibrated to obtain the amyloid fiber-pyrazine aroma compound composite material.

7. The preparation method according to claim 6, characterized in that, The buffer solution is a phosphate buffer solution with a concentration of 10 mmol / L and a pH value of 6.5~7.

5.

8. The preparation method according to claim 6, characterized in that, The mass concentration of amyloid fibers in the mixture is 0.05~0.15%, and the concentration of pyrazine aroma compounds is 0.05~0.10 mmol / L.

9. The preparation method according to claim 6, 7, or 8, characterized in that, The equilibrium was carried out at a temperature of 37°C for 15-20 hours under conditions of darkness.

10. The application of the protein amyloid fiber-pyrazine aroma compound composite material according to any one of claims 1 to 5 or the protein amyloid fiber-pyrazine aroma compound composite material prepared by the preparation method according to any one of claims 6 to 9 in plant-based meat products.

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