Interfacial‐Bulk Synergistic Stabilization of Dual‐Network Emulsion Gels by Protein‐Polysaccharide Ternary Biopolymer System

ABSTRACT

Plant‐based emulsion gels represent sustainable, biocompatible soft materials with transformative potential in food, biomedicine, and advanced manufacturing. However, a long‐standing bottleneck persists in simultaneously achieving long‐term colloidal stability and robust mechanical properties using exclusively natural biopolymers, severely limiting their practical deployment. Herein, we report an interfacial‐bulk synergistic stabilization strategy by constructing a soy protein isolate (SPI)‐konjac glucomannan (KGM)‐κ‐carrageenan (KC) ternary biopolymer system. We systematically elucidate that SPI dominates interfacial adsorption to form elastic interfacial films, while KGM‐KC complexes construct a physically crosslinked bulk network to enhance viscosity and steric hindrance, enabling collaborative stabilization at both interface and bulk phases. Leveraging this synergy, stable high internal phase emulsions are readily fabricated. Furthermore, transglutaminase‐induced covalent crosslinking of SPI interpenetrates with the KGM‐KC physical network, yielding a robust dual‐crosslinked emulsion gel with significantly enhanced mechanical strength, water‐holding capacity, and freeze‐thaw stability. Notably, the optimized gels exhibit excellent 3D printability with high shape fidelity, demonstrating great potential for personalized food manufacturing and bioactive delivery. This work provides fundamental insights into protein‐polysaccharide synergistic interactions and establishes an effective strategy for engineering high‐performance all‐natural soft materials.

An interfacial‐bulk synergistic stabilization strategy is proposed for fabricating dual‐network emulsion gels using protein‐polysaccharide ternary biopolymers. The complementary roles of proteins and polysaccharides enable simultaneous interfacial stabilization and bulk‐phase reinforcement, endowing emulsion gels with superior mechanical strength, excellent stability and high‐precision 3D printability, providing a green platform for all‐natural soft materials.

1. Introduction

Emulsion gels are multi‐structured soft materials composed of dispersed oil droplets, the oil–water interface, and a three‐dimensional (3D) gel network in the continuous aqueous phase. These systems integrate the high loading capacity of emulsions with the tunable mechanical properties of hydrogels, serving as a versatile platform for developing advanced soft materials and investigating fundamental phenomena in colloid and interface science, including interfacial adsorption kinetics, steric and electrostatic stabilization, and multi‐scale gelation mechanisms [1, 2]. In recent years, plant‐based emulsion gels have attracted increasing attention as sustainable and biocompatible alternatives to conventional systems that rely on synthetic surfactants or petroleum‐derived polymers. Driven by demands for environmentally friendly and renewable colloidal materials, researchers have turned to natural macromolecules, particularly plant‐derived proteins and polysaccharides, as stabilizers and gelling agents [3, 4]. These biopolymers are advantageous due to their inherent amphiphilicity (proteins) or chain‐entanglement capability (polysaccharides), as well as their biodegradability and low environmental impact [3, 4, 5, 6, 7]. However, constructing fully plant‐based emulsion gels that simultaneously achieve long‐term colloidal stability and robust mechanical properties remains a persistent challenge in colloid and interface science. This challenge primarily arises from the complementary yet incomplete functionalities of proteins and polysaccharides when used individually or in binary combinations.

Proteins such as soy protein isolate (SPI) are excellent interfacial stabilizers owing to their amphiphilicity and ability to form viscoelastic films at the oil–water interface [5, 6]. However, most protein‐based emulsion gels frequently suffer from limited long‐term stability and weak mechanical strength under extreme conditions due to the denaturation of proteins, which compromises their processability and potential applications [7, 8, 9]. In contrast, polysaccharides such as konjac glucomannan (KGM) and κ‐carrageenan (KC) demonstrate unique advantages in constructing 3D gel networks with high water‐holding capacity and mechanical reinforcement [10, 11]. Notably, binary polysaccharide systems further enhance structural stability and mechanical properties of gel networks through synergistic effects such as intermolecular helical aggregation and chain entanglement [12, 13, 14]. The flexible long chains of KGM and the double‐helix structure of KC form a “rigid‐flexible” complementary structure through intermolecular forces (e.g., hydrogen bonding), yielding semi‐interpenetrating gel networks with enhanced mechanical strength and stability at optimal ratios [15, 16]. This KGM‐KC combination outperforms many other common binary polysaccharide blends, such as KC/locust bean gum, by providing superior gel strength, elasticity, and water‐holding capacity at relatively lower concentrations [17, 18]. Nevertheless, the strong hydrophilicity of polysaccharides severely limits their interfacial activity, thereby restricting their emulsifying capability in stabilizing emulsions [19, 20, 21].

To overcome these complementary limitations, the development of protein‐polysaccharide complex systems has been recognized as a powerful strategy for achieving dual‐phase stabilization of emulsion gels: rapid interfacial adsorption by proteins coupled with bulk‐phase gelation by polysaccharides [22, 23]. While preliminary studies have demonstrated the synergistic effect between proteins‐polysaccharide ternary biopolymer systems in regulating the structure and functions of emulsion gels, most studies have primarily focused on the final macroscopic properties of emulsion gels [24, 25, 26]. The underlying multi‐scale mechanisms, including interfacial adsorption kinetics, interfacial dilational rheology, coupling between interfacial films and bulk networks, and the resulting structure–property relationships of emulsion gels remain largely unexplored.

In this study, we selected SPI as the primary interfacial stabilizer owing to its inherent amphiphilicity and excellent interfacial activity. By integrating SPI with KGM and KC, a ternary biopolymer system (SPI‐KGM‐KC) was developed, enabling the successful preparation of high internal phase emulsions and emulsion gels with tunable properties. The incorporation of transglutaminase (TGase) induced the formation of a covalently crosslinked SPI network that interpenetrated with the physically crosslinked KGM‐KC network to construct a robust dual‐network gel structure. Through multi‐scale in situ characterization, this work systematically elucidates the interfacial‐bulk synergistic stabilization mechanisms in SPI‐KGM‐KC ternary biopolymer systems. Notably, the resulting dual‐network emulsion gels exhibited superior stability, mechanical strength, and environmental resilience. As a proof‐of‐concept for soft‐matter fabrication, these emulsion gels also demonstrated excellent extrusion adaptability and shape fidelity in 3D printing. These findings provide a conceptually novel and generalizable framework for the rational design of high‐performance biopolymer‐based emulsion gels, advancing fundamental understanding of interfacial phenomena and network reinforcement in complex colloidal systems.

2. Experimental Section

2.1. Materials

KGM (purity≥86%) was provided by Yizhi Konjac Biotechnology Co., Ltd. (Hubei, China). Kappa‐carrageenan (KC, purity≥90%) was obtained from Lvxin Food Co., Ltd. (Fujian, China). SPI and soybean oil were purchased from Wanderful Biotechnology Co., Ltd. (Shandong, China). Soybean oil was bought from the local supermarket in Shenzhen. Transglutaminase (TGase) was provided by Dongsheng Biotechnology Co., Ltd. (Jiangsu, China). Sudan Red III, perylene, Rhodamine B, and Rhodamine B isothiocyanate were purchased from Macklin Biochemical Technology Co., Ltd. (Shanghai, China). Calcofluor white stain solution was purchased from Yuanye Biotechnology Co., Ltd. (Shanghai, China). All other chemicals were of analytical grade and used without further purification.

2.2. Preparation of Emulsifier Aqueous Solutions

For the preparation of sole emulsifier solutions, powders of SPI, KGM, and KC were individually dispersed in deionized water at 25°C, followed by mechanical stirring for 20 min in an 80°C water bath to obtain aqueous solutions. For the preparation of binary (KGM‐KC) and ternary (SPI‐KGM‐KC) emulsifier solutions, the powders were co‐dispersed in deionized water, and the remaining steps were identical. The prepared aqueous solutions included: SPI at concentrations of 1 wt%, 2 wt%, and 3 wt%; KGM and KC individually at 0.1 wt%, 0.2 wt%, and 0.3 wt%; KGM‐KC binary mixtures at 0.005 wt%‐0.005 wt%, 0.1 wt%‐0.1 wt%, 0.2 wt%‐0.2 wt%, and 0.3wt%‐0.3 wt%; and SPI‐KGM‐KC ternary mixtures at 1 wt%‐0.05 wt%‐0.05 wt%, 2 wt%‐0.05 wt%‐0.05 wt%, 3 wt%‐0.05 wt%‐0.05 wt%, 1 wt%‐0.1 wt%‐0.1 wt%, and 1 wt%‐0.15 wt%‐0.15 wt%.

2.3. Preparation of Oil‐in‐Water (o/w) Emulsions and Emulsion Gels

For the preparation of o/w emulsions, soybean oil and the biopolymer aqueous solutions prepared in Section 2.2 were preheated to 50°C. The oil phase was mixed with the aqueous phase and subsequently homogenized for 3 min at 8000 rpm using a high‐speed homogenizer (Model XHF‐DY, Ningbo Scientz Biotechnology Co., Ltd., China). To investigate the effect of oil phase fraction, o/w emulsions were prepared at oil‐to‐water mass ratios of 1:2, 1:1, 2:1, 3:1, and 4:1.

For the preparation of emulsion gels, higher concentrations of biopolymer aqueous solutions (10 wt% SPI; 0.5 wt% KGM‐0.5 wt% KC; 10 wt% SPI‐0.5 wt% KGM‐0.5 wt% KC) were prepared following the procedure in Section 2.2. After adding soybean oil at the predetermined ratio, the mixture was homogenized at 10,000 rpm for 10 min using a high‐speed homogenizer. To induce covalent crosslinking of SPI networks, 1 wt% TGase (based on total emulsion mass) was added immediately after homogenization. The resulting emulsion was poured into cylindrical molds, incubated in a 50°C water bath for 1 h to promote crosslinking, and then transferred to an 85°C water bath for 20 min to inactivate TGase. The formulations of as‐prepared emulsion gels are listed in Table S1.

2.4. Dynamic Light Scattering (DLS) and Zeta Potential Measurements

The hydrodynamic radius distribution of different biopolymers was determined by DLS using a dual‐angle, dual‐detector laser light scattering spectrometer (model ALV/CGS‐3, ALV GmbH, Germany) equipped with a multi‐tau digital correlator (model ALV‐7004). The measurements were conducted at a scattering angle of 30°. The intensity autocorrelation function G⁽²⁾(q,t) was analyzed using the relationship Γ = Dq² to obtain the diffusion coefficient distribution G(D). The hydrodynamic radius distribution f(R_h) of different biopolymers was subsequently calculated using the Stokes–Einstein equation as follows:

$$R_h=\frac{k_{B}T}{6\pi \eta D}$$ (1)

where R_h is the hydrodynamic radius, k_B is the Boltzmann constant, T is the absolute temperature, η is the solvent viscosity, and D is the diffusion coefficient.

The zeta potential of different biopolymers was measured at 25°C using a zeta potential and nanoparticle size analyzer (NanoBrook 90Plus PALS, Brookhaven, USA). Prior to the measurement, the samples were highly diluted with deionized water. The refractive index of the dispersion medium was set to 1.33.

2.5. Fourier Transform Infrared (FTIR) Spectroscopy

The FTIR spectra of different biopolymers were recorded using a Fourier transform infrared spectrometer (Thermo Scientific iS50, USA). The spectra were scanned in the range of 4000–400 cm^{− 1} with a resolution of 2 cm^{− 1}.

2.6. Optical Microscopy and Confocal Laser Scanning Microscopy (CLSM)

The morphology of the emulsion droplets was observed using an optical microscope (Nikon Upright Microscope Ci‐L plus, Japan). The oil phase was stained with Sudan Red III prior to observation. Then, a 5 µL aliquot of the stained emulsion was placed on a glass slide and covered with a coverslip. Optical microscopy images were captured using a 10x objective lens. Droplet size and distribution were statistically analyzed with ImageJ software.

CLSM (Nikon AX, Nikon, Japan) was used to visualize the microstructure of emulsion droplets and interfaces according to the reported method with modifications [27]. SPI and polysaccharides were stained with Rhodamine B isothiocyanate and Calcofluor White, respectively, under magnetic stirring in the dark for 4 h, followed by dialysis at least five times to remove free dye. The oil and aqueous phases were stained with perylene and Rhodamine B, respectively. CLSM images were acquired under excitation wavelengths of 408 and 543 nm using a 20× objective lens.

2.7. Evaluation of Emulsion Stability by Centrifugation

To evaluate the stability of as‐prepared emulsions, emulsion samples were placed in centrifuge tubes and centrifuged at 8,000 g for 10 min using a high‐speed centrifuge (Heraeus Multifuge X1R, Thermo Fisher Scientific, Germany). After centrifugation, three distinct phases were observed in the tube: the aqueous phase at the bottom, the emulsion phase in the middle, and the oil phase on the top. The emulsion phase was carefully removed and weighed. Centrifugation stability was calculated as follows:

$$\mathrm{Centrifugation}\mathrm{stability}\left(\%\right)=\frac{m_2}{m_1}\times 100\%$$ (2)

where m ₁ is the initial mass (g) of the emulsion and m ₂ is the mass (g) of the remaining emulsified phase after centrifugation.

2.8. Measurements of Mechanical Properties

The mechanical properties of emulsions were measured at 25°C using a HAAKE MARS 40 rheometer (Thermo Fisher Scientific, Germany) equipped with a 35 mm parallel‐plate geometry (working gap = 1 mm). The apparent viscosity of emulsions was determined via steady‐state shear tests with shear rates ranging from 0.1 s^{− 1} to 100 s^{− 1}. In the linear viscoelastic range, frequency sweeps were performed at a fixed strain of 1% over frequencies from 0.01 Hz to 100 Hz, and strain sweeps were conducted at a fixed frequency of 1 Hz with strains varying from 0.01% to 100% [28].

2.9. Measurements of Contact Angle, Dynamic Interfacial Tension (IFT), and Interfacial Dilational Rheology

The contact angle, IFT, and interfacial dilational rheology of different biopolymers were measured by an all‐purpose contact angle measuring & contour analysis system (OCA‐15EC, Dataphysics) and analyzed with SCA 20 software. For contact angle measurements, the sessile drop method was used. Before measurement, a small amount of the biopolymer aqueous solution was dripped on the surface of a glass slide and dried at room temperature for 24 h to form a thin film. Subsequently, a 3 µL droplet of deionized water was deposited on the film in the air. The droplet profile was captured immediately, and the water contact angle can be automatically calculated by SCA 20 software. All contact angle measurements were repeated 3 times on the same substrates [29].

For IFT and interfacial dilational rheology measurements, the pendant drop method was applied. Typically, a 15 µL water droplet was injected at 15 µL/s from the syringe and suspended in the soybean oil for continuous measurement. The dynamic IFT was monitored in real time for at least 1200 s at 50 °C and determined by fitting the droplet profile to the Young‐Laplace equation [30]. Based on the recorded instantaneous IFT and surface areas, the interfacial dilational rheology was measured according to the method reported by Guan et al. with modifications [31]. The changes in interfacial complex modulus of different biphasic systems were measured over a 3800 s period at a fixed frequency of 0.1 Hz and an amplitude of 10%. After reaching equilibrium, a frequency sweep test was conducted at a fixed amplitude of 10%, with the frequency ranging from 0.005 Hz to 1.0 Hz. An amplitude sweep test was conducted at a fixed frequency of 0.1 Hz, with amplitudes of 5%, 10%, 15%, 20%, 25%, and 30%.

2.10. Food 3D Printing of Emulsion Gels

Before printing, 3D models were constructed using SolidWorks software. The 3D printing of emulsion gels was performed at 25°C with a FOODBOT‐D1 printer (Hangzhou Shiyin Co. Ltd., China). The printing parameters were set as follows: nozzle height of 1 mm, nozzle diameter of 1.20 mm, infill density of 100%, and printing speed of 15 mm/s. Printing performance was evaluated using the following metrics:

$$\mathrm{Printing}\mathrm{accuracy}\mathrm{of}\mathrm{height}\left(\%\right)=\left(1-\frac{\left|H_0-H\right|}{H}\right)\times 100\%$$ (3)

$$\mathrm{Printing}\mathrm{accuracy}\mathrm{of}\mathrm{length}\left(\%\right)=\left(1-\frac{\left|L_0-L\right|}{L}\right)\times 100\%$$ (4)

$$\mathrm{Printing}\mathrm{stability}\left(\%\right)=\frac{H_1}{H_0}\times 100\%$$ (5)

where H₀ and L₀ are the measured height and length of the printed sample, respectively; H and L are the designed height and length of the model, respectively; and H₁ is the height of the sample after standing for 12 h after printing.

2.11. Characterization of Emulsion Gels

The characterization of emulsion gels included the microstructure visualization, macroscopic rheological measurements, texture profile analysis, creep and recovery tests, water holding capacity determination, and freeze‐thaw stability evaluation. The details can be found in the Supporting Information.

3. Results and Discussion

3.1. Preparation of Plant‐Based o/w Emulsions and HIPEs

3.1.1. Physicochemical Properties and Emulsifying Capability of Proteins and Polysaccharides

Prior to emulsification, the physicochemical properties of the protein and polysaccharides were characterized. Zeta potential measurements indicate that SPI, KC, and KGM all carried negative charges under neutral conditions, with KC exhibiting the strongest negative charge (approximately −55 mV) (Figure S1). DLS analysis reveals that proteins and polysaccharides existed in an aggregation state in aqueous solutions (Figure S2). Specifically, SPI formed polydisperse aggregates with hydrodynamic radii centering around 10, 92, and 364 nm. In contrast, KC and KGM exhibited relatively uniform hydrodynamic size. The predominant radius of KGM was around 46 nm with a broader distribution likely arising from its flexible chains adopting various conformations in solution, whereas KC showed a larger and more uniform size (∼ 228 nm), attributable to its rigid double‐helical structure formed upon cooling. As shown in Figure 1, both SPI and the polysaccharides alone were able to form o/w emulsions. For SPI‐stabilized emulsions, the average droplet size was approximately 60 µm at 1 wt% SPI. When the SPI concentration increased to 2 wt%, the droplet size reduced to 54 ± 14 µm with a narrower size distribution, indicating greater interfacial coverage by additional SPI (Figure 1a,b). However, further increasing SPI concentration to 3% did not lead to a notable reduction in droplet size, suggesting saturation of the oil–water interface, with excess SPI remaining dispersed in the aqueous phase without contributing to stabilization

FIGURE 1.

Optical microscopy images and droplet size distributions of o/w emulsions stabilized by different concentrations of (a, b) SPI, (c, d) KC, (e, f) KGM, and (g, h) KGM‐KC.

In comparison, due to the larger hydrodynamic radius, the o/w emulsion stabilized by 0.1 wt% KC showed a larger droplet size (76 ± 28 µm), while KGM at the same concentration formed significantly smaller droplets (40 ± 13 µm) (Figure 1c–f). Notably, droplet sizes of emulsions stabilized by either KGM or KC showed minimal concentration dependence across 0.1–0.3 wt%. When KGM and KC were used simultaneously, the resulting emulsions exhibited smaller droplet sizes and narrower size distribution compared to single‐polysaccharide systems, indicating a synergistic effect between KGM and KC in emulsion stabilization (Figure 1g,h). However, at higher concentrations of KGM and KC, the droplet size distribution widened slightly, likely due to the increased continuous phase viscosity hindering uniform emulsification.

3.1.2. Stabilization and Properties of o/w Emulsions by SPI‐KGM‐KC Ternary Biopolymers

We then investigated the synergistic effects of protein‐polysaccharide ternary emulsifiers on emulsion stabilization. As shown in Figure 2a, SPI, KGM‐KC, and SPI‐KGM‐KC were used as emulsifiers, respectively, to stabilize o/w emulsions under the same emulsifying conditions. CLSM images indicate that SPI can adsorb at oil–water interfaces after emulsification, forming a thin layer on droplet surfaces and preventing droplet coalescence (Figure 2b). In contrast, KGM‐KC exhibited negligible adsorption at oil–water interfaces (Figure 2c). Instead, it is suspected that the rigid double‐helix structures of KC entangle with flexible KGM chains, thereby constructing a 3D network in the continuous aqueous phase. This binary polysaccharide‐based network enhanced emulsion stability by providing steric hindrance to restrict droplet movement and collision. Compared with single or binary emulsifiers, SPI‐KGM‐KC ternary emulsifiers demonstrated a synergistic effect in emulsion stabilization. It is clearly observed that SPI mainly adsorbed at droplet surfaces to form a protective layer, while KGM‐KC primarily formed a physically crosslinked network filling the interstitial spaces between droplets (Figure 2d). Additionally, the emulsions stabilized by SPI‐KGM‐KC exhibited significantly reduced droplet sizes. These results confirmed the synergistic effect of the ternary biopolymer system on interfacial and bulk‐phase stabilization of emulsions.

FIGURE 2.

(a) Schematic illustration of the preparation process and microstructure of o/w emulsions stabilized by SPI, KGM‐KC, and SPI‐KGM‐KC, respectively. Optical microscopy and CLSM images of emulsions stabilized by (b) SPI, (c) KGM‐KC, and (d) SPI‐KGM‐KC. SPI was labeled with Rhodamine B isothiocyanate (red), and KGM‐KC was stained with Calcofluor White (blue). (e) Droplet size distribution and (f) centrifugation stability of emulsions stabilized by SPI‐KGM‐KC at different concentrations and mass ratios. Different letters indicate significant differences at p < 0.05 among samples.

To further elucidate the distinct roles of SPI and KGM‐KC within the ternary biopolymer system, the effects of emulsifier concentration on emulsion properties were investigated. As shown in Figures 2e and S3, when the KGM‐KC concentration was kept constant, increasing the SPI concentration from 1 to 3 wt% significantly reduced the average droplet size from 18 µm to 8 µm and improved the uniformity of droplet sizes. This result indicates that higher SPI concentrations facilitate the stabilization of larger oil–water interfaces. In contrast, variations in KGM‐KC concentration had negligible effects on average droplet sizes, while higher KGM‐KC concentrations slightly increased the polydispersity of droplet sizes. These observations suggest that KGM‐KC primarily contributes to emulsion stability by increasing continuous phase viscosity, thereby hindering the movement of droplets rather than directly participating in interfacial adsorption.

Since emulsion stability is critical for potential applications, we subsequently evaluated the storage and centrifugation stability of emulsions stabilized by various emulsifiers. The results show that all emulsions underwent varying degrees of coalescence after 24 h of storage at room temperature (Figure S4). Especially, the emulsion stabilized solely by SPI or KGM‐KC showed significant increases in droplet size, highlighting the limitations of single proteins or binary polysaccharides for long‐term emulsion stability. In comparison, emulsions stabilized by SPI‐KGM‐KC exhibited smaller changes in droplet size and better storage stability, demonstrating that the combination of SPI with KGM‐KC effectively prevented droplet coalescence. The improved stability of the ternary biopolymer system can be attributed to two complementary mechanisms: (i) the strong negative charge of KGM‐KC in the continuous phase provides electrostatic repulsion between similarly charged SPI‐stabilized droplets, and (ii) hydrogen bonding interactions between SPI and KGM‐KC enhance the anchoring effect of droplets to the continuous bulk network (Figure S5), thereby restricting droplet movement and preventing potential droplet collision. Furthermore, a higher concentration of KGM‐KC was more effective than that of SPI in improving the storage stability of emulsions. When KGM‐KC concentration increased to 0.15 wt%‐0.15 wt% in the ternary biopolymer system, the droplet size remained almost unchanged after 24 h of storage. The emulsion stability was also evaluated by centrifugation tests. Figure 2f indicates that the centrifugation stability of emulsions increased significantly at higher KGM‐KC concentrations, with almost no phase separation observed at 0.15 wt% KGM‐0.15 wt% KC. In comparison, though increasing the SPI concentration effectively reduced the average droplet size, the improvement of emulsion centrifugation stability was relatively limited. These consistent results demonstrate that SPI plays a dominant role in reducing droplet sizes, whereas the presence of KGM‐KC primarily contributes to the long‐term emulsion stability. This interfacial‐bulk division of labor represents a new dual‐phase synergistic stabilization mechanism that overcomes the intrinsic trade‐off between interfacial activity and bulk mechanical reinforcement in single or binary biopolymer systems.

Figure 3 shows the rheological properties of as‐prepared emulsions. All emulsions displayed typical shear‐thinning behavior over the shear rate range of 0.1–100 s^{− 1}, with viscosity decreasing significantly as shear rate increased (Figure 3a,b). It was observed that higher concentrations of SPI or KGM‐KC led to increased emulsion viscosity. The viscosity increase at higher SPI concentrations can be primarily attributed to smaller droplet sizes [32], whereas the increase was more pronounced with higher KGM‐KC concentrations due to the formation of a physically crosslinked network in the bulk phase. Consistent with the viscosity trends, strain sweep tests showed that both storage modulus (G′) and loss modulus (G″) of emulsions increased with rising concentrations of SPI or KGM‐KC (Figure 3c,d). Specifically, G′ exceeded G″ at 3 wt% SPI or 0.15 wt% KGM‐0.15 wt% KC, indicating a transition from a liquid‐like emulsion state to an elastic and solid‐like emulsion state. Notably, increasing the KGM‐KC concentration led to a more significant rise in G′ and the critical strain (γ_c) compared with SPI, suggesting that polysaccharides are more effective in enhancing the structural stability of emulsions and their resistance to external deformation. The structural stability enhancement by KGM‐KC can be attributed to their thickening effect. For example, an aqueous solution containing 1 wt% KGM exhibited a viscosity as high as 100,000 Pa·s at low shear rates, far exceeding that of a solution containing SPI at the same concentration (Figure S6).

FIGURE 3.

Effect of SPI and KGM‐KC concentration variations on the (a, b) viscosity as a function of shear rate and (c, d) rheological properties as a function of oscillation strain for as‐prepared emulsions.

3.1.3. Interfacial Properties of Ternary Biopolymer Systems

To elucidate the underlying stabilization mechanisms of emulsions, the adsorption kinetics and interfacial behavior of SPI, KGM‐KC, and SPI‐KGM‐KC at the oil–water interface were further investigated. Water contact angle measurements revealed that KGM‐KC is highly hydrophilic, with a contact angle of approximately 30°, whereas SPI exhibited a significantly higher contact angle (∼80°), indicating stronger hydrophobicity (Figure 4a). The dynamic IFT measurements indicated that SPI rapidly reduced the IFT to a lower value, whereas KGM‐KC only slightly reduced the IFT during long‐time measurement (Figure 4b,c), suggesting SPI exhibited higher interfacial activity than KGM‐KC. These results are consistent with previous findings that particles with contact angles close to 90° are more likely to adsorb at the oil–water interface, thereby more effectively lowering the IFT [33]. Notably, a slight decrease in the IFT for the pristine oil–water interface was observed even without protein or polysaccharides, likely due to the presence of surface‐active substances (e.g., fatty acids and phospholipids) in soybean oil (Figure S7) [34, 35]. These surface‐active substances in the oil phase may also contribute to emulsion formation and stabilization. In ternary emulsifier systems, increasing the SPI concentration from 0.1 to 0.3 wt% accelerated the initial IFT reduction (0–100 s) and resulted in lower IFT during 1000 s measurement as more SPI diffused and adsorbed at the interface during the same period (Figure 4b). This result aligns well with the observed decrease in emulsion droplet size as SPI concentration increased (Figure S3). However, with the fixed SPI concentration (0.1 wt%), the effect of KGM‐KC concentration on IFT was overall negligible. Specifically, the IFT of the ternary biopolymer system containing 0.1 wt% SPI and 0.005 wt% KGM‐0.005 wt% KC was only marginally lower than the system containing 0.1 wt% SPI. Furthermore, increasing the KGM‐KC concentration led to a relatively slower initial IFT reduction and a higher equilibrium IFT. This may be attributed to the increased aqueous‐phase viscosity and competitive adsorption induced by KGM‐KC, which hindered the diffusion of SPI from the bulk phase to the interface and accordingly reduced the interfacial coverage of SPI.

FIGURE 4.

(a) Water contact angles of SPI and KGM‐KC. Different letters indicate significant differences at p < 0.05 among samples. (b, c) Dynamic IFT and (d‐f) interfacial dilational rheology of pendant droplets covered by SPI, KGM‐KC, and SPI‐KGM‐KC as functions of (d) time, (e) frequency, and (f) amplitude.

As the stability of emulsions primarily relies on the mechanical properties of the interfacial film, the effect of different emulsifiers on the interfacial dilatational rheology was evaluated. Figure 4d shows the time‐dependent evolution of the complex modulus (E*) of oil–water interfaces covered by various emulsifiers. The results indicate E* increased gradually with adsorption time for all biphasic systems, reflecting increasing interfacial coverage. Specifically, the E* for the KGM‐KC‐covered interface was extremely low, indicating that polysaccharides form interfacial films with limited mechanical strength. In contrast, the SPI‐covered interface exhibited the highest E*, demonstrating that protein adsorption can impart the interface with superior robustness. Notably, incorporating KGM‐KC into SPI‐covered interfaces resulted in a reduction in E*, with further reductions in E* as KGM‐KC concentration increased, whereas increasing SPI concentration in the ternary biopolymer system promoted an enhancement in E*. This suggests that the interfacial elasticity is predominantly governed by the adsorbed SPI layer, whereas the introduction of KGM‐KC disrupted SPI packing structures at the interface to some extent. Lissajous plots were used to further analyze the viscoelasticity of interfacial films during the measurement (Figure S8). At the early adsorption stage (360 s), the Lissajous plots for all interfaces were elliptical in both compression and expansion processes due to the low interfacial coverage, suggesting typical viscoelastic behavior of interfaces. After adsorption for 3480 s, except for the interface solely covered by KGM‐KC, all interfaces containing SPI evolved to elongated shapes with a higher surface pressure, showing strain hardening and pronounced elastic response during droplet deformation, indicating the formation of more elastic interfacial films [36]. Among all systems, droplets covered by 0.1 wt% SPI‐0.005 wt% KGM‐0.005 wt% KC exhibited the best compression resistance and shape recovery compared to The those covered by single or binary emulsifiers, suggesting that small amounts of KGM‐KC impart enhanced flexibility and toughness to the interfacial film. However, this synergistic effect diminished at higher concentrations of either SPI or KGM‐KC, highlighting the presence of a critical optimal ratio for SPI and KGM‐KC to achieve effective interfacial synergy. Based on this result, we hypothesize that low level of KGM‐KC can intercalate between SPI molecules at the interface, preventing overly dense and brittle structures and forming a balanced rigid‐flexible composite network. The elastic response of various interfaces was also supported by pendant drop appearance evolution during deformation. As shown in Figure S9, no interfacial jamming or buckling occurred on KGM‐KC‐covered droplet surfaces even under substantial compression, whereas pronounced wrinkles can be observed on SPI‐covered droplet surfaces under minor compression, indicating high interfacial coverage of SPI (> 82.93%). Similar wrinkling was observed on ternary emulsifier‐covered droplet surfaces, confirming that the interface is predominantly occupied by SPI.

The dynamic mechanical response of interfacial films was further investigated via frequency and amplitude sweeps. The results show that E* for KGM‐KC‐covered droplet surfaces increased significantly with frequency, which is indicative of typical fluid‐like interfacial behavior (Figure 4e). In contrast, all SPI‐covered surfaces, regardless of KGM‐KC presence, exhibited weak frequency dependence and solid‐like interfacial behavior, indicating the formation of rigid and elastic interfacial films by SPI [37]. As amplitude increased from 5% to 25%, all SPI‐containing interfacial films showed strain softening (Figure 4f), suggesting irreversible structural breakdown under large deformation [30]. In comparison, KGM‐KC‐covered surfaces exhibited lower E* with no clear amplitude dependence, likely due to the rearrangement of flexible polysaccharide chains accommodating deformation via segment slippage [38]. These findings establish quantitative guidelines for tuning interfacial viscoelasticity in ternary biopolymer systems, offering broader implications for the design of stable complex fluids and bio‐based interfaces.

3.1.4. Stabilization and Mechanical Properties of HIPEs

Building on the excellent emulsifying capability and interfacial properties of SPI‐KGM‐KC ternary emulsifiers, we further explored their potential for stabilizing o/w HIPEs. As shown in Figures 5a and S10, o/w emulsions stabilized by SPI‐KGM‐KC ternary emulsifiers exhibited outstanding stability across various oil‐water ratios. When the oil fraction exceeded 74 vol%, stable HIPEs were successfully prepared. CLSM images revealed that droplets in the HIPEs were closely packed, deforming from regular spherical shapes into irregular polyhedral shapes, with SPI adsorbed at droplet surfaces to prevent coalescence (Figure 5a). The successful preparation of HIPEs can be attributed to the synergistic effect of SPI‐KGM‐KC, which effectively encapsulates a high volume of oil droplets while forming a stable 3D gel network. In contrast, single or binary emulsifiers (SPI or KGM‐KC) only formed o/w emulsions at lower oil‐to‐water ratios with poorer stability, exhibiting obvious creaming or phase separation after storage for 24 h (Figure S10).

FIGURE 5.

(a) Macroscopic appearance and corresponding microstructure of the HIPE with an oil‐to‐water ratio of 4:1 stabilized by SPI‐KGM‐KC. (b) Viscosity as a function of shear rate and (c) rheological properties as a function of angular frequency for ternary emulsifier‐stabilized emulsions at different oil‐to‐water ratios. (d) Rheological properties for the HIPE with an oil‐to‐water ratio of 4:1 as a function of oscillation strain.

The microstructure and rheological behavior of ternary emulsifier‐stabilized HIPEs were further investigated as a function of oil‐to‐water ratio. As the oil phase ratio increased, droplet number rose accordingly, and the droplet size became less uniform (Figure S11). At an oil‐to‐water ratio of 3:1, obvious droplet deformation occurred due to the spatial confinement of droplets and increasing packing pressure, representing a typical microstructural feature of HIPEs [39]. The dense packing of droplets significantly enhanced the inter‐droplet interactions and friction, leading to elevated apparent viscosity of HIPEs compared to emulsions with lower oil fractions (Figure 5b). Additionally, when the oil‐to‐water ratio was 1:1, G′ and G″ of the emulsion were nearly comparable over a broad frequency range. However, once the oil‐to‐water ratio exceeded 2:1, G′ became significantly higher than G″, indicating that emulsions underwent a transition from a fluid‐like state to a predominantly elastic and solid‐like state (Figure 5c). As the oil phase ratio further increased, both G′ and the critical strain (γ_c) rose markedly (Figures 5d and S12). These results demonstrate that higher internal phase fractions enable the emulsion to withstand greater strain while maintaining structural integrity.

3.2. Preparation and Property Regulation of Emulsion Gels

3.2.1. Enzymatic Crosslinking‐induced Emulsion Gel Formation

TGase is known to catalyze the formation of ε‐(γ‐glutamoyl)lysine isopeptide bonds between the lysine and glutamine residues abundant in SPI [40, 41]. Since SPI was dispersed both in the continuous phase and adsorbed at droplet surfaces, the introduction of TGase could achieve covalent crosslinking among SPI molecules, thus leading to covalently crosslinked networks. Meanwhile, KGM‐KC forms a non‐covalent physically crosslinked network through chain entanglement. Figure 6a illustrates the preparation process as well as the impact of internal phase fraction on the microstructure of the emulsion gels in the presence of TGase. We suspect that the covalently crosslinked gel network formed at low internal phase fractions is primarily derived from SPI in the continuous phase. In comparison, at higher internal phase fractions, dense droplet packing drives the formation of a more compact covalently crosslinked gel network, which is constructed not only by free SPI in the continuous phase but also via interactions between closely packed droplets. CLSM images provide direct visualization of the dual‐network structure in the emulsion gel (Figure S13). Specifically, in the absence of TGase, SPI and SPI‐covered droplets were well dispersed within the continuous phase. The introduction of TGase induced the formation of a crosslinked network comprising SPI and SPI‐covered droplets, which interpenetrated with the KGM‐KC crosslinked network. The synergistic coupling between the covalently crosslinked SPI network and the physically crosslinked KGM‐KC network leads to emulsion gels with significantly enhanced mechanical properties and stability.

FIGURE 6.

(a) Schematic illustration of the preparation process of emulsion gels at varying oil‐water ratios in the presence of TGase. (b) Macroscopic appearance and (c) corresponding SEM images of prepared emulsion gels with different formulations. (d) Temperature sweep and frequency sweep curves of prepared emulsion gels with different formulations.

We then investigated the effects of composition and internal phase fraction on the macroscopic appearances and microstructures of emulsion gels. As shown in Figure 6b, emulsion gels stabilized solely by SPI or KGM‐KC at a low oil‐water ratio (1:3) exhibited poor structural integrity and were prone to collapse. In contrast, the utilization of SPI‐KGM‐KC ternary emulsifiers significantly improved the self‐supporting capacity of the resulting emulsion gels. Moreover, the introduction of TGase further reinforced gel structural integrity through enzymatic crosslinking, which endowed the emulsion gels with a smoother and more compact surface morphology. Notably, despite the high initial mechanical strength of emulsion gels derived from HIPE, the accelerated crosslinking within the concentrated system induced the formation of rigid gel aggregates before molding, ultimately leading to heterogeneous macroscopic appearances. Corresponding SEM images provided further insights into the internal microstructural differences among emulsion gels with different formulations (Figure 6c). Specifically, the TGase‐crosslinked emulsion gel stabilized solely by SPI featured a rough, porous, and heterogeneous network structure after freeze‐drying. In contrast, lyophilized KGM‐KC‐stabilized emulsion gels formed a lamellar structure, which contributes to enhancing the toughness of polysaccharide‐based gels [42]. For emulsion gels co‐stabilized by SPI‐KGM‐KC ternary emulsifiers, a homogeneous network with uniform pore size distribution was formed. Upon the addition of TGase in the ternary biopolymer system, the interconnected pores of lyophilized emulsion gels almost disappeared due to the synergistic effect between covalently crosslinked SPI networks and physically crosslinked KGM‐KC networks in the continuous phase, which significantly thickened the pore walls and prevented droplet rupture during freeze‐drying. At an oil‐to‐water ratio of 3:1, emulsion gels without TGase exhibited notably increased porosity and interconnected pore structures after freeze‐drying. This phenomenon is associated with the dense packing of droplets in HIPEs. Similarly, the introduction of TGase into this system promoted the formation of a denser gel network with fewer interconnected pores.

Given that G′ is positively correlated with the degree of intermolecular crosslinking during gelation [43], we further elucidated the gelation mechanism of emulsion gels via temperature sweep tests (Figure 6d). The entire gelation process can be divided into three stages. Before testing, all TGase‐containing samples exhibited higher initial G′ than TGase‐free systems, indicating that TGase had already induced preliminary covalent crosslinking of SPI at room temperature. During the isothermal incubation at 50°C for 1 h (stage 1), G′ gradually increased in all samples, with a more pronounced elevation in TGase‐added samples, demonstrating efficient formation of SPI networks at 50°C. Interestingly, in the early phase of this period, the sample stabilized solely by SPI without TGase showed a faster G′ increment from a much lower initial level compared to the one containing TGase. This unique phenomenon can be attributed to heat‐induced SPI aggregation driven by non‐covalent interactions, though the resulting network lacks structural robustness due to the absence of covalent crosslinks. The incubation temperature was then ramped to 85°C and subsequently held for 20 min to inactivate TGase (stage 2). In this stage, samples without TGase exhibited an initial G′ decrease followed by recovery, whereas TGase‐containing samples were less affected by temperature increases. Notably, the sample stabilized solely by KGM‐KC underwent severe structural collapse during 85°C holding due to the disruption of chain entanglement. In contrast, G′ continued to increase in all SPI‐containing samples as thermal denaturation further strengthened the crosslinked SPI network by forming protein aggregation [44]. These results suggest that stage 2 primarily disrupted weak non‐covalent interactions (e.g., hydrogen bonds), while the covalently crosslinked SPI network maintained the structural integrity of emulsion gels. During cooling to 25°C (stage 3), the gel network gradually solidified. Decreasing temperature promoted the reconstruction and reinforcement of non‐covalent interactions between various components, driving further network densification. However, G′ decreased during cooling in HIPE‐derived emulsion gels, likely due to internal stress arising from mismatched thermal contraction rates between the continuous gel phase and dispersed oil droplets, which disrupted local network continuity and reduced modulus.

Frequency sweep results demonstrated that, except for the emulsion gels stabilized by SPI or KGM‐KC without TGase, all other samples exhibited weak frequency dependence of G′ across the tested frequency range (0.1–10 Hz). The G′ of emulsion gels stabilized by SPI‐KGM‐KC was significantly higher than those stabilized by single or binary emulsifiers, confirming the synergistic effect between the SPI and KGM‐KC. Additionally, at the same oil‐water ratio, TGase‐containing samples showed higher G′, indicating that enzymatic crosslinking effectively strengthened the gel network. Compared to emulsion gels at a low oil‐to‐water ratio (1:3), HIPE‐derived emulsion gels also exhibited higher G′, which is attributed to the combined effects of dense droplet packing and the dual‐crosslinked gel network, collectively constructing a more robust gel structure with enhanced mechanical properties.

3.2.2. Textural Properties, Dynamic Mechanical Properties, and Freeze‐Thaw Stability of Emulsion Gels

The textural properties of emulsion gels are critical for developing their potential applications in functional foods [45]. To mimic the oral mastication process, texture profile analysis was performed to evaluate the hardness, springiness, cohesiveness, and chewiness of different emulsion gels. Figure 7a shows that the emulsion gel stabilized by KGM‐KC exhibited the lowest hardness. By incorporating SPI, the hardness of ternary emulsifier‐stabilized emulsion gels was significantly enhanced. Furthermore, either the incorporation of TGase or increasing the oil‐phase ratio to 3:1 further enhanced gel hardness substantially. Among all samples, HIPE‐derived emulsion gels stabilized by SPI‐KGM‐KC with TGase exhibited the highest hardness, which is consistent with the frequency sweep results, primarily attributable to the synergistic effect between the dual‐network structure, denser droplet packing, and higher oil fraction. In contrast, Figure 7b indicates that the springiness of emulsion gels was predominantly governed by polysaccharides. The incorporation of SPI into KGM‐KC‐stabilized emulsion gels reduced their springiness, suggesting that SPI enhanced the overall plasticity of the gel network. However, TGase‐induced covalent crosslinking of SPI reversed this trend and contributed to improved springiness. Notably, there was no significant difference in cohesiveness among the various emulsion gels. The emulsion gel stabilized solely by SPI with TGase exhibited the highest cohesiveness (Figure 7c), which is attributed to the highly uniform and tightly connected covalent protein network formed via enzymatic crosslinking. Additionally, both enzymatic crosslinking and a higher oil‐water ratio led to a significant improvement in the chewiness of emulsion gels. This trend is consistent with that of hardness, as chewiness is predominantly governed by the initial hardness of the emulsion gel [46].

FIGURE 7.

(a–d) Textural properties of emulsion gels, including (a) hardness, (b) springiness, (c) cohesiveness, and (d) chewiness. (e, f) Dynamic mechanical properties of emulsion gels, including (e) creep‐recovery curves and (f) creep compliance curves. (g) Water holding capacity and (h) freeze‐thaw stability of emulsion gels. Different letters indicate significant differences at p < 0.05 among samples.

The deformation behavior and recovery capacity of emulsion gels under sustained stress were further evaluated via creep recovery tests. As shown in Figure 7e, during the creep phase (0–2 min) under constant stress, the strain of all samples increased with time. Emulsion gels stabilized solely by physical entanglement of KGM‐KC exhibited the highest flexibility, showing the maximum strain and the lowest recovery rate after stress removal. In comparison, SPI‐stabilized emulsion gels with TGase displayed significantly reduced strain under the same stress, and their recovery rate was higher than that of KGM‐KC‐stabilized emulsion gels, indicating that covalently crosslinked networks are more prone to resist stress compared to physically crosslinked networks. The synergistic effect between SPI and KGM‐KC endowed the ternary emulsifier‐stabilized emulsion gels with enhanced resistance to sustained stress and improved recovery rates. Additionally, a higher oil‐water ratio further enhanced the deformation resistance and recovery performance of emulsion gels. Notably, ternary emulsifier‐stabilized emulsion gels with both dual‐crosslinked networks and high oil‐water ratios exhibited the minimal creep strain and highest recovery rate, confirming the formation of a stable 3D network with structural integrity. The creep compliance of the emulsion gels showed a trend consistent with their creep‐recovery profiles. KGM‐KC‐stabilized emulsion gels exhibited the highest creep compliance due to the relatively weak gel structures, whereas the incorporation of SPI‐KGM‐KC, a higher oil‐to‐water ratio, and TGase‐induced crosslinking effectively reduced the creep compliance of as‐prepared emulsion gels (Figure 7f).

Water‐holding capacity (WHC) is a critical indicator for evaluating the application potential of gel‐based foods. As shown in Figure 7g, all prepared emulsion gels exhibited a WHC exceeding 80%, demonstrating excellent water retention performance. Among them, KGM‐KC‐stabilized emulsion gels exhibited the optimal WHC, owing to the abundant hydrophilic groups on their molecules and the dense physically entangled gel networks, which effectively entrapped water molecules within the 3D gel structure via hydration [47]. For emulsion gels at an oil‐to‐water ratio of 1:3, the effect of TGase on the WHC was negligible. However, for HIPE‐derived gels, the introduction of TGase was particularly critical, as it worked in concert with the high internal phase fraction and dense droplet packing to reinforce both interfaces and the bulk network to resist structural collapse and water exudation, thereby achieving a higher WHC. Figure 7h shows the freeze‐thaw stability of emulsion gels. After multiple freeze‐thaw cycles, KGM‐KC‐stabilized emulsion gels maintained excellent stability with a low syneresis, which may be attributed to their unique lamellar network structure. This structure can inhibit the migration and growth of ice crystals, while the elastic network can alleviate the mechanical stress generated by ice crystal growth through reversible deformation, thereby mitigating structural damage [48, 49]. Furthermore, the covalently crosslinked network formed via TGase catalysis further enhanced the freeze‐thaw stability of emulsion gels. Notably, HIPE‐derived emulsion gels exhibited significantly higher syneresis because of dense droplet packing and thinner gel networks. As a result, ice crystal growth during freezing was more likely to rupture droplet surfaces, leading to irreversible phase separation.

3.3. Applicability of SPI‐KGM‐KC‐Stabilized Emulsion Gels in 3D Printing

3D printing has emerged as a powerful fabrication technique for complex soft‐matter architectures in various fields [50, 51]. Leveraging the high mechanical strength and shear‐thinning behavior of the SPI‐KGM‐KC‐stabilized emulsion gels, we further evaluated the applicability of such emulsion gel systems as printable inks. As shown in Figure 8a, the letters “S”, “Z”, and “U” were successfully printed onto a substrate. Owing to the relatively low mechanical strength, the emulsion gel without TGase exhibited post‐extrusion flowability that compromised line precision and resulted in blurred edges after printing. In contrast, TGase addition enhanced structural support, yielding printed letters with clearer and sharper edges. Although the emulsion gel at a high oil‐to‐water ratio (3:1) demonstrated excellent self‐supporting properties, its excessively high viscosity limited shear‐thinning during printing, causing unstable jetting and the roughest edges of printed letters. In comparison, the emulsion gel at a low oil‐to‐water ratio (1:3) with TGase achieved the highest printing accuracy in both planar dimensions and vertical height (Figures 8c,d). After standing in the air for 12 h post‐printing, all printed letters experienced varying degrees of height collapse and oil exudation. Among them, the TGase‐crosslinked emulsion gel at an oil‐to‐water ratio of 1:3 retained the most complete shape, demonstrating superior stability (Figure 8e). However, the HIPE‐derived emulsion gel exhibited significant oil exudation owing to compromised emulsion stability caused by the reduced continuous aqueous phase. To identify the underlying cause, additional experiments were conducted by comparing sealed and non‐sealed conditions (Figure S14). The sealed samples exhibited almost no visible shrinkage and maintained nearly 100% weight retention after 6 h, whereas the non‐sealed samples showed significant structural collapse and weight loss. These results verify that the collapse in height and structural deformation observed herein originate predominantly from moisture evaporation, rather than internal stress generated by sustained gelation reactions. Furthermore, the stability of printed emulsion gels in aqueous environments was assessed via immersion in deionized water for 24 h. As shown in Figure 8f, only the TGase‐crosslinked emulsion gel largely preserved its original structure, whereas the other two samples underwent obvious structural collapse and droplet disassembly. These results underscore the robustness of the dual‐network structure and illustrate the broader utility of interfacial‐bulk synergistic stabilization in engineering printable emulsion gels for advanced materials applications.

FIGURE 8.

(a) Appearance of 3D‐printed emulsion gel letters. (b) Shape retention performance of 3D‐printed letters after 12 h of air storage. (c) Printing accuracy comparison of the actual height versus the designed height of printed letters. (d) Printing accuracy comparison of the actual length versus the designed length of printed letters. (e) Post‐printing stability of printed letters. (f) Integrity of the emulsion gel before and after 24 h immersion in deionized water. Different letters indicate significant differences at p < 0.05 among samples.

4. Conclusion

In summary, we propose an interfacial‐bulk synergistic stabilization strategy using SPI‐KGM‐KC ternary biopolymers to fabricate oil‐in‐water emulsions with tunable structures and properties. Multi‐scale in situ characterization reveals that SPI dominates interfacial tension reduction and elastic film formation, while KGM‐KC enhances stability via increased continuous‐phase viscosity and steric hindrance. This dual‐phase synergistic effect not only significantly improves long‐term emulsion stability but also enables the facile preparation of stable HIPEs. Furthermore, the introduction of TGase induces covalent crosslinking of SPI, which interpenetrates with the physically crosslinked KGM‐KC network to construct a robust dual‐network emulsion gel.

This dual‐phase synergistic mechanism overcomes a long‐standing challenge in the field that single or binary biopolymer systems cannot simultaneously achieve high emulsification efficiency and superior mechanical properties [5, 11, 22, 23]. Additionally, conventional strategies typically require complex modification to improve the interfacial activity of single or binary biopolymers, which is both time‐ and energy‐consuming. Although protein‐polysaccharide ternary biopolymer systems have recently emerged as promising stabilizers for emulsion gels, offering improved stability and mechanical strength relative to traditional single or binary systems, most prior studies have focused predominantly on the macroscopic performance of emulsion gels for specific food applications [24, 25, 26]. Critically, no systematic investigation has been conducted to clarify the synergistic mechanism among ternary biopolymers in emulsion stabilization and gel formation. To fill this critical knowledge gap, we offer an in‐depth understanding of the interfacial adsorption kinetics and interfacial rheology of SPI and KGM‐KC via in situ interfacial characterization, thereby enabling precise regulation of the properties of emulsion gels.

Compared to the emulsion gels with comparable protein and polysaccharide concentrations [52, 53], the resulting dual‐network emulsion gels exhibit superior mechanical strength, excellent water‐holding capacity, enhanced freeze‐thaw stability, and better 3D printing accuracy/shape fidelity. These improvements stem from the unique interfacial‐bulk synergistic stabilization and interpenetrating dual‐network structure, which provide more robust reinforcement than previously reported single or binary crosslinking strategies, thus enabling high‐precision 3D printing with outstanding shape fidelity. These findings not only provide fundamental insights into the interfacial‐bulk synergistic interactions of protein‐polysaccharide ternary systems, but also offer a facile, green, and versatile platform for engineering high‐performance all‐natural soft materials. This strategy can be readily extended to other plant‐derived protein‐polysaccharide combinations, opening broad application prospects in functional foods, cosmetics, and biomedical delivery systems.

Author Contributions

Yue Zheng and Wendi Liang: Investigation, Methodology, Formal analysis, Visualization, Data curation, Writing – original draft. Teng Cao: Investigation, Methodology. Hao Chang: Methodology, Formal analysis. Zhou Liu and Tiantian Kong: Methodology, Validation. Xin Guan: Supervision, Conceptualization, Validation, Project administration, Writing – original draft, Writing – review & editing. Lianwei Li: Supervision, Conceptualization, Validation, Project administration, Writing – review & editing.

Funding

The National Natural Science Foundation of China (22322303), the Natural Science Foundation for Distinguished Young Scholars of Guangdong Province (2023B1515020001), the Guangdong Basic and Applied Basic Research Foundation (2026A1515012441), the Shenzhen Science and Technology Program (JCYJ20250604182413018, QNXMC20250701094307010).

Conflicts of Interest

The authors declare no competing financial interest.

Supporting information

Supporting File: smll74728‐sup‐0001‐SuppMat.pdf.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.