Oleogels Based on Chickpea Protein Fractions–Xanthan Gum Complexes: Preparation and Characterization

TL;DR

Researchers studied how different chickpea protein fractions mixed with xanthan gum form stable oil gels, finding that glutelin-based gels performed best.

Contribution

The study reveals the structural and interfacial properties of chickpea protein fractions in oleogel formation, offering new insights for food formulation.

Findings

GLU-XG complexes formed the smallest particles and most stable emulsions with high centrifugal stability.

GLU-XG-based oleogels showed the highest oil-binding capacity and storage modulus.

Fluorescence spectroscopy showed GLU-XG had higher surface hydrophobicity and conformational flexibility.

Abstract

This study investigated the mechanism by which different fractions of chickpea protein influenced the formation of oleogels. Total chickpea protein (CPP, 0.5 wt%), chickpea albumin (ALB, 0.5 wt%), globulin (GLO, 0.5 wt%), and glutelin (GLU, 0.5 wt%) were separately used as oleogelators by combining with xanthan gum (XG, 0.5 wt%) at pH 7 to construct soybean oil-based oleogels via the emulsion-templated method. Particle size measurement revealed that the GLU-XG (526 nm) exhibited the smallest particle size compared to CPP-XG (605 nm), ALB-XG (642 nm), and GLO-XG (819 nm). The four complexes exhibited increasing surface hydrophobicity and conformational flexibility (as revealed by fluorescence spectroscopy) in the order of GLO-XG < ALB-XG < CPP-XG < GLU-XG. Compared with other complexes, the higher surface hydrophobicity, smaller particle size, and more flexible structure of GLU-XG conferred a superior surface activity. Consequently, the fabricated emulsion demonstrated a smaller droplet size (13.91 μm) and enhanced centrifugal stability (94.64%). The confocal laser scanning microscope images confirmed that the oleogel based on GLU-XG exhibited the most uniform and densest network, leading to the highest oil-binding capacity (98.7%) and storage/loss modulus, followed by those based on CPP-XG (97.2%), ALB-XG (95.6%), and GLO-XG (93.9%). This research provides a theoretical basis for using chickpea protein in oleogel formulations and enhances understanding of the structural and interfacial properties of these protein fractions.

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1. Introduction

Solid fats confer flavors, textures, and mouthfeel to food products, occupying a significant position in the modern edible oils industry. However, traditional solid fats are rich in saturated or trans fatty acids. Excessive consumption increases health risks, including cardiovascular disease and obesity [1]. Consequently, numerous countries have imposed restrictions on their content in foods [2,3]. In recent years, structuring unsaturated plant oils into edible soft solids as a healthier alternative to solid fats has attracted extensive attention [4]. Oleogels are usually constructed by oleogelators, in which liquid oils are trapped in three-dimensional networks. As a novel and promising solid fat substitute with zero trans fatty acids and low saturated fatty acids, oleogels have potential applications in the food industry. Low-molecular-weight oleogelators (LMWOGs) are generally used to prepare oleogels by the direct method [5]. However, oil deterioration caused by high-temperature heating and the limited addition of LMWOGs greatly restricts the use of oleogels in foods. In contrast, the indirect method utilizing biopolymers (such as proteins and polysaccharides) for oleogel preparation has attracted extensive attention from researchers, with the emulsion templating method emerging as one of the most mature techniques for converting liquid oils into oleogels [6]. The principle of the emulsion templating method involves preparing O/W emulsions using biopolymers, subsequently removing the water phase, and shearing the mixture to form oleogels. Consequently, the properties of these biopolymers not only determine the stability and microstructure of the intermediate emulsion state but also influence the final oleogel’s properties through variations in interfacial behavior and network-forming capabilities.

Chickpea (Cicer arietinum L.), as the second-most globally consumed bean, boasts a rich protein content (25–31%) [7]. Numerous studies have indicated that chickpea protein exhibits a high bioavailability, low allergenicity, and excellent functional properties (such as emulsifying and foaming capabilities) [8]. Based on the emulsifying capacity of chickpea protein, its isolate has been employed in combination with polysaccharides in a previous study to formulate Pickering emulsion systems [9]. Chickpea protein comprises three principal fractions, namely saline–soluble globulin (53–60%), acid/alkali–soluble glutelin (19–25%), and water–soluble albumin (8–12%) [10,11]. Different protein fractions exhibit intrinsic differences in molecular weight, molecular structure, and surface hydrophobicity, potentially leading to distinct behaviors in solubility, surface activity, and intermolecular interaction capacity. For instance, the compact structure of globulin typically reduces molecular flexibility, leading to relatively lower emulsifying properties [12]. Albumin exhibited greater solubility than other proteins due to its lower molecular weight [13].

Variations in the molecular properties of different fractions (such as molecular size, conformational flexibility, surface hydrophobicity, and interaction propensity) systematically influence their adsorption kinetics at the oil–water interface, the structural–rheological properties of the interfacial film, and the subsequent formation and strengthening of networks during dehydration [14]. These effects ultimately manifest as significant differences in emulsion stability and macroscopic texture of the oleogels. Generally, fractions with rapid solubility and high molecular flexibility can preferentially and quickly adsorb at the interface, effectively reducing interfacial tension. In contrast, fractions with larger molecular weight and compact structures may form stronger cohesive forces within the interfacial adsorption layer or the continuous phase through hydrophobic interactions, disulfide bonds, and other mechanisms, thereby enhancing the mechanical strength of the interfacial film and the overall rigidity of the gel network. For instance, in the soy protein system, 7S globulin exhibits superior emulsifying properties to 11S globulin owing to its superior solubility and flexibility [14]. Meanwhile, 11S globulin, being more prone to thermal aggregation and forming more disulfide bonds, tends to produce gels that are firmer and more viscoelastic [15]. Similarly, studies on chickpea protein have shown that its different fractions exhibit distinct emulsifying and foaming properties [7].

To date, total plant/animal proteins have been widely used as naturally derived oleogelators, such as canola protein [16], soybean protein [17], and gelatin [18]. Some researchers have also employed a specific single protein fraction, such as zein [19]. Furthermore, they have focused on comparing differences between proteins from various sources in oleogel formation. For instance, Annika et al. [20] investigated the key factors influencing oleogel strength using five plant and animal proteins. Athira et al. [21] compared the properties of pea- and faba bean-based oleogels and assessed their potential as a baking ingredient. However, whether examining total proteins or specific fractions, current work predominantly remains at the level of macroscopic characterization of oleogel properties and often treats total protein as a functional whole. Consequently, there is a lack of systematic comparisons between different fractions of the same protein. Moreover, it remains unclear which properties of protein fractions determine the characteristics of the resulting oleogels, or how these properties influence protein network formation. We hypothesize that protein fractions with differing molecular properties may affect oleogel structure formation due to their distinct interfacial characteristics. As an anionic polysaccharide with a multibranch linear structure, Xanthan gum (XG) exhibits high viscosity. XG can be applied to form stable oleogels in combination with other oleogelators or structuring agents [18,22]. We hypothesize that chickpea protein–XG complexes might also prepare oleogels with higher gel strength. Additionally, soybean oil was chosen as the base oil because it is rich in polyunsaturated fatty acids (linoleic acid content > 50%) and is one of the most commonly consumed edible oils worldwide [23].

In this work, total chickpea protein and its three major fractions (albumin, globulin, and glutelin) were chosen to prepare oleogels via the emulsion-templated method by combining with XG. The structures and properties of these oleogels were compared. This study offers significant insights into the development of chickpea protein fraction-based oleogels.

2. Materials and Methods

2.1. Materials

Chickpea flour was purchased from Mulei Farmer Brothers Agricultural Development Co., Ltd. (Mu Lei County, China). Soybean oil was purchased from a local market. XG was obtained from Shanghai Yuanye Bio-Technology Co., Ltd. (Shanghai, China). All other reagents were of analytical purity.

2.2. Extraction of the Total Protein and the Three Protein Fractions

The total chickpea protein (CPP) was isolated using a previous method with minor modifications [24]. In brief, the chickpea flour was defatted with hexane. Then the obtained sediment was mixed with deionized water at a solid-to-liquid ratio of 1:10 (w/v) and pH 10.5, under continuous stirring for 2 h at ambient temperature. After centrifugation at 8000× g (Allegra 64R Centrifuge, Beckman, CA, USA) for 20 min, the supernatant was adjusted to pH 4.5 and then centrifuged (8000× g, 20 min). The obtained precipitates were dissolved in deionized water at pH 7.0 and dialyzed in deionized water at 4 °C for 48 h, followed by freeze-drying at −80 °C for 48 h (Freezone^®^6 Plus, Labconco, Kansas City, KS, USA). In addition, according to the Osborne extraction method [25], the three chickpea protein fractions, albumin (ALB), globulin (GLO), and glutelin (GLU), were separately extracted from the above total protein using deionized water, 5% NaCl, and 0.1 mol/L NaOH with a solid-to-liquid ratio of 1:10 (w/v). The following steps were similar to those of the total protein extraction. The protein content of the four freeze-dried powders was measured using the Bradford protein assay [26]; the detailed procedure is provided in the Supplementary Materials.

2.3. Preparation of Protein–XG Complexes

The protein powders (CPP, ALB, GLO, GLU) were dissolved in deionized water to prepare 0.5 wt% solutions. Then, XG powder was added to each solution to prepare protein–XG complexes with a final XG concentration of 0.5 wt%, and the pH of each complex solution was adjusted to 7. The obtained complexes were named CPP-X, ALB-X, GLO-X, and GLU-X, respectively. Some parts of complex solution samples were stored at 4 °C, and the others were freeze-dried for further analysis.

2.4. Characterization of Proteins and Protein–XG Complexes

2.4.1. Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) Analysis

The subunit distributions of freeze-dried CPP, ALB, GLO, and GLU were analyzed using SDS-PAGE [27]. Concentrations of stacking and separating gels were set at 5% and 12.5%, respectively. Each sample was mixed with SDS-reducing buffer containing 2% SDS (w/v). After heating at 100 °C for 10 min, the samples were cooled and then centrifuged at 1200× g for 5 min. The supernatant from each sample (7.0 μL) was then loaded into each well. The electrophoresis voltage was maintained at 180 V.

2.4.2. Surface Hydrophobicity

The samples (four proteins and their corresponding complexes) were dissolved in phosphate buffer (0.01 M, pH 7.0) and diluted to final protein concentrations of 0.01, 0.05, 0.1, 0.25, and 0.5 mg/mL, respectively. The fluorescence intensity of these samples was measured using 1-Anilinonaphthalene-8-sulfonic acid (ANS) probe. A 4 mL sample was mixed with 20 μL of 8 mM ANS solution, and the reaction was carried out in the dark for 10 min. A fluorescence spectrophotometer (F-7000, Hitachi, Tokyo, Japan) was used to measure fluorescence intensity at an excitation wavelength of 390 nm and an emission wavelength of 470 nm. For each sample, a scatter plot was created with protein concentration (mg/mL) on the X-axis and the corresponding relative fluorescence intensity (a.u.) on the Y-axis. A linear regression analysis was performed on this plot. The equation of the best-fit line was obtained. The hydrophobicity intensity (H_0_) was defined as the slope of the linear fit of fluorescence intensity to sample concentration.

2.4.3. Fluorescence Spectroscopy

The fluorescence intensities of the four proteins, protein–XG complexes, and XG were determined using a fluorescence spectrophotometer. The emission spectra were recorded from 290 to 450 nm with an excitation wavelength of 280 nm.

2.4.4. Particle Size and ζ-Potential

The particle sizes and ζ-potentials of the four proteins, protein–XG complexes, and XG were measured using a Nano ZS Zetasizer (Nano-ZS 2000, Malvern Instruments Ltd., Worcestershire, UK) at 25 °C. Before analysis, each solution was diluted 100-fold with deionized water (pH 7.0).

2.4.5. Fourier Transform Infrared Spectroscopy (FTIR)

The infrared spectra of the four proteins, protein–XG complexes, and XG were recorded using a Fourier transform infrared spectrometer (Spectrum 100, Perkin-Elmer, Warrington, UK). Briefly, 2.0 mg of the lyophilized powders were mixed with 200 mg dry KBr, separately, and then the mixtures were pressed into pellets. The spectral data were collected from 4000 to 600 cm^−1^ at a resolution of 4 cm^−1^.

2.5. Preparation of Protein–XG Complexes Stabilized Emulsions and Oleogels

The oil phase (soybean oil) and the aqueous phase (protein–XG complex solutions) were mixed at a 4:6 (v/v) ratio, and the mixture was homogenized with a homogenizer (PhD Technology LLC, Saint Paul, MN, USA) at 9000 rpm for 40 s to obtain emulsions. Some of the prepared emulsions were used for the following characterization measurements. The other parts were pre-frozen at −80 °C for 24 h, then lyophilized at −80 °C for 48 h under a vacuum of less than 10 Pa. Finally, the oleogels were obtained by simple stirring. The emulsions made by CPP-X, ALB-X, GLO-X, and GLU-X were named CPP-XE, ALB-XE, GLO-XE, and GLU-XE, respectively. The obtained oleogels were correspondingly named CPP-XO, ALB-XO, GLO-XO, and GLU-XO, respectively.

2.6. Characteristics of the Emulsions

2.6.1. Droplet Size

The droplet size of the emulsions was determined with a laser particle size analyzer (LS230, Beckman Coulter, Inc., Brea, CA, USA). The refractive indices were 1.470 and 1.333 for soybean oil and water, respectively. The volume–weight diameter (D [4,3]) was used to represent the results.

2.6.2. Centrifugal Stability

Centrifugal stability of the emulsions was measured according to the previously described method, with some modifications [28]. A total of 200 μL of the emulsion was diluted with 3 mL deionized water, and the OD value at 500 nm (A_0_) was measured using an ultraviolet–visible spectrometer (TU-1901, Beijing Puxi General Instrument Co., Ltd., Beijing, China); the sample was centrifuged at 2000× g for 10 min. Subsequently, the residual emulsion was pipetted from the bottom of the centrifuge tube, and its OD was measured as described above (A_1_). The centrifugal stability was calculated according to the following equation:

$$[eqn]$$

2.6.3. Emulsion Microstructures

The microstructures of the emulsions were analyzed with an upright fluorescence microscope (DM4 B, Leica Microsystems, Wetzlar, Germany) after staining the emulsions for 30 min with a mixed dye solution (1 mg/mL Nile red and 1 mg/mL fluorescein isothiocyanate (FITC)). The excitation wavelengths of 568 nm (Nile red) and 488 nm (FITC) were used for sample observation.

2.7. Characterization of the Oleogels

2.7.1. Oil Binding Capacity (OBC)

The OBC was determined via centrifugation [29]. Briefly, 1.0 g of the oleogel was placed in a 1.5 mL tube and centrifuged (7000× g, 20 min, 25 °C). The released liquid oil was removed. OBC was calculated using the equation:

$$[eqn]$$

where m is the mass of the residual oleogel after centrifugation (g), and m_0_ is the mass of the initial oleogel (g).

Stability tests for oleogels were conducted to measure the OBC of the samples at 0, 3, 6, 9, 12, 15, and 20 d during storage at 20 °C.

2.7.2. Confocal Laser Scanning Microscope (CLSM)

Following the staining procedure detailed in Section 2.6.3, fresh oleogels were imaged using a CLSM (AXR NSPARC, Nikon Corporation, Tokyo, Japan).

2.7.3. Rheological Properties

The rheological properties of the fresh oleogel were determined using a rotational rheometer (Discovery HR-2, TA Instruments, West Sussex, UK). Measurements were performed at 25 °C using a steel parallel plate with a 40 mm diameter, with a 1 mm gap between the plates. At a constant frequency of 1 Hz, amplitude sweeps were conducted over a strain range of 0.01% to 100% to determine the linear viscoelastic region (LVR). Frequency sweeps were performed at a fixed strain of 0.1% with angular frequency ranging from 0.1 to 100 rad/s. The storage modulus (G′) and loss modulus (G″) were measured as functions of frequency. The apparent viscosity of the oleogels was evaluated through steady-shear flow tests over a shear rate range of 0.1 to 100 s^−1^. In addition, the oleogels underwent three-interval thixotropy tests (3-ITT) under alternating shear rates (0.1 s^−1^, 10 s^−1^, and 0.1 s^−1^) to evaluate their recovery abilities. Temperature sweeps (frequency = 1 Hz, strain: 0.1%) were conducted at the temperature variation of 25–85 °C with a rate of 5 °C/min.

2.7.4. Differential Scanning Calorimetry (DSC)

The thermal behaviors of the fresh oleogels were investigated using a differential scanning calorimeter (DSC214, NETZSCH, Selb, Germany). Oleogels (0.0050 g) were placed in aluminum pans and subjected to a heating process from 25 °C to 150 °C. N_2_ atmosphere (40 mL/min) was used as the purging gas. Soybean oil was used as a control.

2.8. Statistical Analysis

Each experiment was repeated at least in triplicate. The data were analyzed using a one-way ANOVA with Duncan’s test in SPSS 26.0 software (IBM Corporation, Armonk, NY, USA) to determine statistically significant differences (p < 0.05). The data were presented as the mean ± standard deviation.

3. Results and Discussion

3.1. Protein Contents and SDS-PAGE of the Chickpea Protein Fractions

The protein contents of CPP, ALB, GLO, and GLU were 93.12 ± 1.54%, 89.16 ± 2.36%, 92.45 ± 1.14%, and 93.40 ± 2.39%, respectively.

As shown in Figure 1A, CPP exhibited polypeptide bands within 13–100 kDa, and the major bands were located at 20, 24, 40, 51, and 65 kDa. The 20 and 51 kDa bands correspond to subunits of the 7S globulin, while the 24 and 40 kDa bands correspond to subunits of the 11S globulin [25]. ALB displayed a similar polypeptide band distribution to that of CPP. However, its major bands at 11–17 kDa were clearly observed, indicating that ALB mainly comprised low-molecular-weight polypeptides, consistent with its water-soluble protein fraction. Additionally, both GLO and GLU displayed the major bands at 20, 24, and 40 kDa. Although the two protein fractions had similar polypeptide band patterns, different extraction conditions (especially pH) might lead to structural differences between them, further affecting their functional characteristics [7]. The observed molecular weight distributions resembled the patterns reported by Gao et al. [7], but showed slight differences from those reported by Chang et al. [25]. These differences were likely attributable to the different source materials. The results demonstrated that the four proteins were successfully separated and extracted.

3.2. Surface Hydrophobicity of the Chickpea Protein Fractions

The H_0_ of proteins is one of the key parameters for evaluating the exposure of surface hydrophobic groups, which are directly related to protein interface properties [10]. As shown in Figure 1B, GLU presented the highest H_0_, followed by CPP, ALB, and GLO. Generally, a higher H_0_ indicated a greater exposure of surface hydrophobic groups and a higher degree of protein unfolding. This structural change promoted the rapid adsorption of protein molecules at the oil–water interface, resulting in smaller droplets and a denser interfacial coating [30], which in turn yielded a stable oleogel network.

In comparison with that of the single protein, the surface hydrophobicity of the protein–XG complexes significantly decreased with the addition of XG. The reason might be that XG bound to the surface of proteins, thereby inhibiting the binding of ANS fluorescent probes to hydrophobic amino acids [31,32]. Although the addition of XG reduced the surface hydrophobicity of the complexes, its thickening effect was crucial for stabilizing the emulsions and oleogels. Further discussion was displayed in Section 3.6 and Section 3.7. The surface hydrophobicity of the four complexes showed similar changing trends with the single proteins. The H_0_ values of these four complexes were presented as GLO-X < ALB-X < CPP-X < GLU-X.

3.3. Fluorescence Analysis of the Protein Fractions and Their Complexes

At an excitation wavelength of 290 nm, the intrinsic fluorescence spectra of proteins reflect the polar microenvironment of tryptophan residues, enabling further investigation of protein conformational changes and interactions with XG [33]. As shown in Figure 2, the maximum emission peaks (λ_max_) of CPP, ALB, GLO, and GLU were respectively 330.0, 333.1, 325.1, and 337.1 nm, agreeing with the previous results [7]. The larger λ_max_ suggested that the tryptophan residues of proteins might be located in a more polar microenvironment [34]. Simultaneously, the fluorescence intensity gradually decreased in the order of GLO > ALB > CPP > GLU. The higher fluorescence intensity suggested a more compact protein conformation, which could have a more effective preservation of buried tryptophan residues [35]. Additionally, no emission peaks were observed in the XG spectrum. A larger λ_max_ and lower fluorescence intensity indicated that more hydrophobic regions of the proteins were exposed to the polar microenvironment, and that the proteins existed in more unfolded and flexible conformations. Therefore, the predicted order of conformation flexibility was GLU > CPP > ALB > GLO, which correlated with the results of surface hydrophobicity.

Compared with the single proteins, the fluorescence intensity of the corresponding complexes decreased, and the λ_max_ shifted toward shorter wavelengths, consistent with the findings of Niu et al. [36]. The endogenous fluorescence quenching of the complexes might be ascribed to XG binding to proteins, thereby trapping some tryptophan residues within the side chains of XG or within the structural domains of the complexes. In addition, the weak blue shifts in λ_max_ indicated that the complexation with XG reduced the exposure of the proteins’ hydrophobic groups, and the tertiary structures might be slightly compressed [37]. However, the fluorescence intensities and maximum emission peaks (λmax) of the four complexes showed changes similar to those of the single proteins, indicating that the addition of XG did not significantly affect the conformational flexibility of the four proteins. The conformational flexibility of protein molecules significantly affected their emulsification properties. Enhanced conformational flexibility increased the accessibility of hydrophobic groups within the protein, moving to the oil–water interface, thereby lowering interfacial tension and reducing droplet size [38].

3.4. Particle Size and ζ-Potential of Chickpea Protein Fractions and Their Complexes

It has been reported that higher concentrations of XG made firmer structures of oleogels [39], and also a coarser surface of oleogels. As a result of pre-experimental optimizations, a lower XG concentration (0.5 wt%) was used in our study to obtain a smooth gel-like texture. Moreover, the protein–XG complexes, which were made under pH 7 and a protein concentration of 0.5 wt%, could be used to prepare the following stable emulsions and corresponding oleogels (as shown in Figures S1 and S2). Therefore, the protein–XG complexes prepared under the above conditions were selected as materials for the follow-up experiment. As presented in Figure 3A, CPP and the three protein fractions were negatively charged, and their potential values were around −30 mV, which was attributed to the pH (7.0) of the protein suspensions being higher than the isoelectric point of each protein (Figure S3). In addition, XG also carried negative charges (−55.00  ±  1.28 mV), due to the deprotonation of its carboxyl groups at pH 7 [40]. Although both carried a net negative charge and were unlikely to attract each other via electrostatics, the proteins and XG could still form complexes through hydrophobic interactions and hydrogen bonding, as reported by Zhao et al. [41]. A further discussion of the protein–XG interaction forces is displayed in the next section. The potentials of the four protein–XG complexes were similar to that of XG, and significantly higher than those of the corresponding proteins, implying XG coating on each protein surface through intermolecular forces [41]. Furthermore, the absolute values of the ζ-potential for the four complexes exceeded 30 mV, indicating that these complexes could maintain good emulsion stability [42].

As shown in Figure 3B, the mean sizes of the four proteins were within 210–270 nm with no significant differences (p > 0.05), and XG displayed a particle size of 121.0 ± 6.2 nm. However, the mean size of each complex (in the range of 520–820 nm) was obviously bigger than that of the corresponding protein. Simultaneously, the four complexes’ sizes showed significant differences (p < 0.05), and the sequence of gradual increase was GLU-X < CPP-X < ALB-X < GLO-X.

The structure and strength of protein–polysaccharide complexes depend on the physicochemical properties of the biopolymers, such as the charged groups of the proteins, the molecular flexibility of the native proteins (i.e., the ease of the structural unfolding), the chain flexibility, and the charge distributions on the polysaccharide backbones [43] (pp. 225–259). We hypothesized that differences in particle size among the complexes might be due to conformational variation among the four proteins. Surface hydrophobicity and fluorescence spectroscopy results revealed differences in the exposure of hydrophobic groups and the degree of unfolding among the four proteins, indirectly indicating significant variations in their conformational flexibility. Proteins with higher unfolding degrees (e.g., GLU) may exhibit greater conformational freedom due to their flexibility. This allowed it to rearrange its structures and form multi-point non-covalent entanglements with XG [44,45], thereby reducing cross-linking with additional proteins or XG. Consequently, the resulting GLU-X (525.6 ± 6.6 nm) was also smaller than the other complexes. In contrast, proteins with a higher proportion of globular structures (e.g., GLO) exhibited greater rigidity, making them less prone to bending or twisting. Therefore, it might locally interact with XG at a single specific binding point [46,47]. Since XG surfaces contained multiple active binding sites, each XG molecule could combine with multiple protein molecules, leading to protein aggregation through this bridge effect. And complexes of larger size were formed. Ma et al. [48] characterized lupin protein–polysaccharide complexes via atomic force microscopy, which was consistent with our results. Their work revealed that rigid polysaccharides bind proteins through “single-point bridging”, fostering large aggregates. In contrast, flexible chains enabled “multi-point anchoring” due to their conformational adaptability, thereby reducing cross-linking and producing smaller complexes. As for CPP and ALB, their semi-flexible nature likely enabled more extensive binding with XG than that of GLO [46].

In view of the differences in flexibility among these proteins, GLO-X probably formed the most rigid structures, whereas CPP-X and ALB-X structures might be more flexible, with GLU-X potentially exhibiting the most flexible structure.

3.5. Fourier Transform Infrared Spectroscopy of the Protein Fractions and Their Complexes

The molecular interactions between the protein fractions and XG were investigated using FTIR. The spectra of CPP and its complexes are shown in Figure 3C, and the FTIR results for the other samples showed similar trends (Figure S4). CPP and XG separately showed broad peaks around 3301 cm^−1^ and 3404 cm^−1^, which were attributed to O-H stretching vibrations. When CPP combined with XG, the O-H stretching vibration peak shifted to 3339 cm^−1^, indicating the formation of hydrogen bonding interactions within the complexes [49]. The absorption peaks at 2931 cm^−1^ and 2927 cm^−1^ for CPP and XG originated from C-H stretching vibrations, respectively. The peak at 1656 cm^−1^ in CPP (amide I) was caused by the stretching vibration of C=O in the amide bond. The peak at 1633 cm^−1^ in XG was caused by the stretching vibration of C=O in the carboxyl group. In CPP-X, the C=O stretching band was observed at 1652 cm^−1^, which might be attributed to hydrophobic interactions between CPP and XG [50]. The characteristic peak at 1023 cm^−1^ in XG corresponded to the stretching vibration of the C-O bond, which shifted to 1043 cm^−1^ in CPP-X. This shift might be due to the electrostatic interactions between CPP and XG. Although the proteins and XG both carry a high negative charge, a weak electrostatic interaction might occur between positively charged local patches of the protein and negatively charged groups of the anionic polysaccharide [51]. This phenomenon was also referred to as the “charge patch” effect [52].

3.6. Characterization of Emulsions

Emulsion formation is an important step in oleogel preparation in the emulsion-templated method, as the cross-linked interfacial film formed by the oleogelators within the emulsion ultimately constitutes the framework of the oleogel. The four emulsions stabilized by protein–XG complexes exhibited a homogeneous appearance (Figure 4A). After 3 d of storage at 4 °C, the emulsions with XG were stable, whereas those without XG showed phase separation after 3 d (Figure S5). The difference in emulsion stability was attributed to the addition of XG, which resulted in a denser interfacial layer formed by the complexes at the O/W interface. Additionally, the thickening effect of XG also played the key role.

As shown in Figure 4B, the average droplet sizes for the four freshly made emulsions were about 20 μm, and GLO-XE and GLU-XE exhibited the largest and the smallest dimensions, respectively. The interfacial microstructure of the samples is shown in Figure 5, with green and red fluorescent fields representing the protein and oil phases, respectively. The dense layers surrounding the oil droplets indicated the samples were O/W emulsions, and the droplet sizes were consistent with the results illustrated in Figure 4B. Furthermore, the four protein–XG complexes exhibited different uniformity when adsorbing at the oil–water interface. GLU-X had the most uniform adsorption, followed by CPP-X, ALB-X, and GLO-X. These differences are fundamentally due to variations in interfacial adsorption among distinct protein–polysaccharide complexes during emulsification. A good emulsifier requires the following properties: (i) fast adsorption to the interface; (ii) high ability of its structure to unfold at the interface; and (iii) formation of a cohesive and viscoelastic interfacial coating [53]. This process was collectively determined by the key molecular properties of the complexes, such as size, flexibility, and surface hydrophobicity. Research indicated that smaller complexes with higher hydrophobicity diffused and adsorbed more rapidly to the oil–water interface [54]. Meanwhile, greater molecular flexibility promoted the stretching and twisting of the complexes at the interface [55], which not only facilitated the exposure and adsorption of more hydrophobic groups but also enhanced the packing density and mechanical strength of the interfacial coating [31], thereby effectively stabilizing fine droplets. As demonstrated in the study by Huang et al. [56], the enhanced molecular flexibility of facilitated efficient protein-oil interactions promoted rapid interfacial adsorption and yielded minimized droplet dimensions.

The centrifugal stability of the four emulsions was further investigated. GLU-XE exhibited the highest stability, followed by CPP-XE, ALB-XE, and GLO-XE (Figure 4B). According to Stokes’ law [57], centrifugal stability may be influenced by droplet size, dynamic viscosity, and the density difference between the oil and water phases. The observed differences in centrifugal stability in this study might primarily be attributed to variations in emulsion droplet size. Although distinct protein aqueous solutions might induce subtle variations in the viscosity of the continuous phase and in the density difference between the continuous and oil phases, their influence was significantly less pronounced than the effect of droplet size variations. Consequently, the emulsions with smaller dimensions exhibited better centrifugal stability, consistent with previous studies [58]. Moreover, the increased flexibility of the complex, which formed a more compact viscoelastic coating, also inhibited the flocculation and coalescence of droplets [59]. Smaller droplet sizes, more robust interfacial layers, and superior stability were critical for the subsequent oleogel formation. A stable, high-strength emulsion template was considered more conducive to the formation of oleogels with high-strength network structures [60].

3.7. Appearance and Oil Binding Capacity of Oleogels

The appearance of the lyophilized samples is shown in Figure S6. They just needed a simple stirring to form a gel-like state. As shown in Figure 6A, all four oleogels had an appearance and smooth texture similar to those of solid fats and exhibited good inverted, self-supporting characteristics. This smooth texture might be associated with the formation of the β′-crystals (Figure S7) [61]. In contrast, the emulsions stabilized by the four individual proteins or XG failed to form oleogels (Figure S8), indicating that the addition of XG mainly improved the resistance of the gelators to droplet coalescence during freeze-drying. This also illustrated that the formation of protein–XG complexes was key to constructing stable oleogels, and the stable oleogels were intrinsically linked to the stable emulsion structures formed by these protein–XG complexes (as shown in the results of Section 3.6). Figure 6B shows that the four fresh-made oleogels exhibited excellent OBC (93.9–98.7%), and the OBC of each oleogel gradually decreased with storage time. However, GLU-XO presented the slowest decrease rate, followed by CPP-XO, ALB-XO, and GLO-XO. After 20 d of storage, the OBC of the four oleogels decreased to 90.12% (GLU-XO), 81.14% (CPP-XO), 80.48% (ALB-XO), and 75.80% (GLO-XO), respectively. The OBC of oleogels directly reflected the strength of the three-dimensional network formed by the protein–XG complexes [62]. The relationship between the network structure and OBC is further discussed in the following sections.

3.8. Microstructure of Oleogels

In this study, CLSM was used to observe the microstructures of the oleogels. As shown in Figure 7, red fluorescence (representing the oil phase) occupied the entire field of view, and the oil droplets were wrapped in the cross-linked networks of the complexes (labeled by green fluorescence), demonstrating the typical oleogel structure. Moreover, the oleogels stabilized by the different complexes exhibited diverse morphologies of the porous networks. In GLU-XO and CPP-XO, a higher proportion of more regular spherical networks (shown in red circles) was found. However, ALB-XO and GLO-XO were mainly dominated by an irregular network (shown in blue circles). These differences led to a decrease in the density and uniformity of the oleogel network in the order of GLU-XO > CPP-XO > ALB-XO > GLO-XO, which, in turn, significantly impacted the strength and stability of the oleogels [62]. This was primarily attributed to the fact that increased network density and structural uniformity could enhance the oleogels’ resistance to external disruption, maintain their structural integrity, and effectively minimize the free flow of oil [63], which were closely associated with their OBC and viscoelastic properties.

3.9. Mechanism Analysis of the Oleogel Network Formation

As shown in Figure 8, based on the above experiment results, the formation mechanism of the oleogel networks with different morphologies was deduced. GLU, with its flexible structure, formed a small, open complex with XG, whereas GLO, with its structural rigidity, formed a large, rigid complex. During emulsion preparation, complexes with higher surface hydrophobicity, smaller size, and more flexible structure (e.g., GLU-XG) could adsorb uniformly at the oil–water interface, promoting the formation of emulsions with uniform oil droplets and a dense coating [64]. In contrast, the complexes with larger size and more rigid structure (e.g., GLO-XG) led to emulsion formation with non-uniform oil droplets and a fragile interfacial layer [14]. During emulsion lyophilization, water loss drove the increase in capillary force between the oil droplets, which induced the deformation of the interfacial network constructed from the complexes [65,66]. Compared with the stronger interfacial network, the weaker one was more prone to deformation, further resulting in the formation of an irregular and disordered oleogel network structure [29].

3.10. Rheological Behaviors of the Oleogels

The rheological behaviors of the oleogels were directly governed by their internal network structures. The rheological behavior of the four oleogels is shown in Figure 9. The amplitude sweep (Figure 9A) results indicated that, within the LVR (the strain range of 0.01–1%), the G′ of the four oleogels was higher than the G″, demonstrating that all the oleogels had elastic solid-like properties [67]. Beyond 1% strain, the curve of G′ intersected with the one of G″ and the G′ of all the samples decreased, indicating the breakdown of the oleogel structure [55,68].

The frequency sweep results (performed at 0.1% strain) for the oleogels are presented in Figure 9B. The G′ of all samples exhibited weak frequency dependence, and GIU-XO had the highest G′ value (29.1 kPa), followed by CPP-XO (24.3 kPa), ALB-XO (12.8 kPa), and GLO-XO (10.9 kPa), indicating the four oleogels possessed high gel strength, especially GLU-XO. Furthermore, the oleogels exhibited G′ values (10–100 kPa) comparable to those of commercial butter [69], demonstrating their potential for application as solid fat substitutes.

Figure 9C showed that the viscosity of all samples decreased as the shear rate increased, demonstrating typical shear-thinning behaviors. This phenomenon indicated that shearing action partially destroyed the gel structure, thereby reducing flow resistance [70,71]. GLU-XO showed the highest viscosity, attributed to its dense and uniform network. Additionally, CPP-XO, ALB-XO, and GLO-XO demonstrated similar viscosity within the entire shear rate range, illustrating that the three oleogels with different cross-linked structures had a similar capacity to resist the shearing action, which agreed with the previous report [65].

The thixotropic behaviors of the samples are shown in Figure 9D. At a low shear rate (0.1 s^−1^), the viscosity of the four oleogels remained stable with the increase in time. While viscosity decreased markedly at a higher shear rate (10 s^−1^), this was attributed to the high-intensity shearing, which facilitated disruption of the network structure [71]. The viscosity rapidly returned to higher values when the shear rate declined to 0.1 s^−1^, which might be due to the good recovery of intermolecular forces that maintain the oleogel structure [65]. Generally, oleogels with a thixotropic recovery exceeding 70% have excellent thixotropic resilience [72], and the internal network structure can be rapidly rebuilt after the removal of shear stress. Table 1 illustrates that GLU-XO exhibited the highest thixotropic recovery (75.45%), followed by CPP-XO (71.91%), ALB-XO (69.35%), and GLO-XO (65.51%), indicating that GLU-XO, CPP-XO, and ALB-XO (especially GLU-XO) demonstrated potential applications in food processing [70,73].

The temperature sweep was performed to assess the thermal sensitivity of the oleogels. As shown in Figure 9E, G′ exceeded G″ throughout the test, confirming that the solid-like elastic behavior dominated. G′ of the four oleogels remained stable with the temperature rising from 25 °C to 85 °C, indicating good thermal stability. This high thermal stability stood in stark contrast to butter’s melting characteristics [69].

Consequently, the oleogels prepared in this study showed a gel strength comparable to that of commercial butter, while possessing a superior thermal stability absent in butter. Coupled with its composition featuring low saturated fat and trans fatty acids, this system demonstrated potential to replace traditional solid fats in food applications requiring both thermal processing stability and healthy fat formulations.

3.11. Thermal Stability Analysis

Thermal stability is another important property of the oleogels. The thermal stability of the four oleogels was evaluated through DSC analysis. As shown in Figure 9F, like soybean oil, the DSC curves of all the oleogels exhibited a straight line without obvious peaks, illustrating that the four oleogels remained stable in the range of 40–150 °C. This observation aligned with the results of the temperature sweep rheological analysis. Furthermore, the melting temperature was considerably higher than that of traditional oleogels stabilized by LMWOGs [74,75].

4. Conclusions

In this study, the feasibility of constructing oleogels based on chickpea protein fractions–XG complexes through the emulsion-templated method was investigated. The four proteins (CPP, ALB, GLO, and GLU) exhibited distinct molecular conformation and interfacial properties, significantly influencing the network structure of the corresponding oleogels and further affecting their physical properties. Compared with CPP, ALB, and GLO, GLU showed the best performance in structural flexibility, interfacial adsorption, and network formation. Accordingly, the oleogel based on GLU-X presented the highest gel network strength, followed by CPP-XO, ALB-XO, and GLO-XO. Furthermore, the OBC sequences and rheological properties correlated with the oleogel network density. The oleogels prepared in this study exhibited enhanced rheological and mechanical properties, along with favorable thermal stability and low saturated fatty acid content, supporting their potential as substitutes for conventional solid fats. However, it is noteworthy that, while GLU showed superior functional properties, its practical application must be evaluated within a broader context that includes economic feasibility, raw material availability, and nutritional value.