Next Article in Journal
Lactobacillus plantarum M3 Fermentation Enhances Mulberry Juice Antioxidant Capacity: Metabolomic Analysis
Previous Article in Journal
Spatiotemporal Differences of 24-Epibrassinolide Regulating Anthocyanin and Proanthocyanidin Biosynthesis in Vitis vinifera ‘Cabernet Sauvignon’
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Foods
Volume 15
Issue 5
10.3390/foods15050905
foods-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

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

by
Xiaomeng Li
1,2,
Songqi Yang
1,2,
Jingwen Wu
1,2,
Yunan Jin
1,2 and
Xiaohong Mei
1,2,*
1
College of Food Science & Nutritional Engineering, China Agricultural University, Beijing 100083, China
2
Key Laboratory of Safety Assessment of Genetically Modified Organism (Food Safety), Ministry of Agriculture and Rural Affairs of the People’s Republic of China, Beijing 100083, China
*
Author to whom correspondence should be addressed.
Foods 2026, 15(5), 905; https://doi.org/10.3390/foods15050905
Submission received: 12 January 2026 / Revised: 14 February 2026 / Accepted: 3 March 2026 / Published: 5 March 2026
(This article belongs to the Section Food Engineering and Technology)
Browse Figures
Versions Notes

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.

Graphical Abstract

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 (H0) 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 (A0) 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 (A1). The centrifugal stability was calculated according to the following equation:
Centrifugal   stability   %   =   A 1 A 0 × 100

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:
OBC   %   = m m 0 ×   100
where m is the mass of the residual oleogel after centrifugation (g), and m0 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. N2 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 H0 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 H0, followed by CPP, ALB, and GLO. Generally, a higher H0 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 H0 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 (λmax) 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 λmax 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 λmax 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 λmax 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 λmax 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.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/foods15050905/s1, Figure S1: Particle size and zeta potential of complexes at different pH (A) and (B). EAI and ESI of complexes at different pH (C) and (D); Figure S2: Particle size and zeta potential of complexes at different protein concentrations (A) and (B). EAI and ESI of complexes at different protein concentrations (C) and (D); Figure S3: Zeta potential of CPP, ALB, GLO, GLU, and XG at different pH; Figure S4: FTIR spectra of ALB, GLO, GLU, XG and their complexes; Figure S5: Appearance of emulsions stabilized by CPP, ALB, GLO, and GLU stored at 4 °C for 3 d; Figure S6: Appearance of the oleogels and their corresponding lyophilized samples; Figure S7: XRD of oleogels; Figure S8: Products of lyophilized samples stabilized by CPP, ALB, GLO, GLU, and XG.

Author Contributions

Conceptualization, X.M.; software, X.L., S.Y., J.W. and Y.J.; investigation, X.L.; data curation, X.L., S.Y., J.W. and Y.J.; writing—original draft preparation, X.L.; writing—review and editing, X.M. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by Guangdong Provincial Key Laboratory of Nutraceuticals and Functional Foods (2024SZKFJJ07).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The original contributions presented in the study are included in the article/Supplementary Materials; further inquiries can be directed to the corresponding author.

Conflicts of Interest

The authors declare no conflicts of interest.

References

  1. Julibert, A.; Bibiloni, M.D.M.; Tur, J.A. Dietary Fat Intake and Metabolic Syndrome in Adults: A Systematic Review. Nutr. Metab. Cardiovasc. Dis. 2019, 29, 887–905. [Google Scholar] [CrossRef] [Scilit]
  2. Hwang, H.-S. A Critical Review on Structures, Health Effects, Oxidative Stability, and Sensory Properties of Oleogels. Biocatal. Agric. Biotechnol. 2020, 26, 101657. [Google Scholar] [CrossRef] [Scilit]
  3. Adili, L.; Roufegarinejad, L.; Tabibiazar, M.; Hamishehkar, H.; Alizadeh, A. Development and Characterization of Reinforced Ethyl Cellulose Based Oleogel with Adipic Acid: Its Application in Cake and Beef Burger. LWT 2020, 126, 109277. [Google Scholar] [CrossRef] [Scilit]
  4. Ma, Y.; Ye, F.; Chen, J.; Ming, J.; Zhou, C.; Zhao, G.; Lei, L. The Microstructure and Gel Properties of Linseed Oil and Soy Protein Isolate Based-Oleogel Constructed with Highland Barley β-Glucan and Its Application in Luncheon Meat. Food Hydrocoll. 2023, 140, 108666. [Google Scholar] [CrossRef] [Scilit]
  5. Giacomozzi, A.S.; Palla, C.A.; Carrín, M.E.; Martini, S. Physical Properties of Monoglycerides Oleogels Modified by Concentration, Cooling Rate, and High-Intensity Ultrasound. J. Food Sci. 2019, 84, 2549–2561. [Google Scholar] [CrossRef] [Scilit]
  6. Feichtinger, A.; Scholten, E. Preparation of Protein Oleogels: Effect on Structure and Functionality. Foods 2020, 9, 1745. [Google Scholar] [CrossRef] [Scilit]
  7. Gao, Y.; Yin, A.; Feng, J.; Zhao, G.; Pang, J.; Yuan, T.; Zheng, H.; Shi, J. Structures, Functional Properties and Simulated Digestion in Vitro of Chickpea Protein Fractions. Eur. Food Res. Technol. 2025, 251, 3287–3303. [Google Scholar] [CrossRef] [Scilit]
  8. Ye, J.; Shi, N.; Rozi, P.; Kong, L.; Zhou, J.; Yang, H. A Comparative Study of the Structural and Functional Properties of Chickpea Albumin and Globulin Protein Fractions. Food Bioprocess Technol. 2024, 17, 3253–3266. [Google Scholar] [CrossRef] [Scilit]
  9. Liu, Y.; Fan, X.; Yu, X.; Liu, T.; Guo, X.; Zhang, J. Enhancing Curcumin Stability and Bioavailability through Chickpea Protein Isolate–Citrus Pectin Conjugate Emulsions: Targeted Delivery and Gut Microecology Modulation. Int. J. Biol. Macromol. 2025, 300, 140295. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  10. Perović, M.N.; Tomičić, Z.M.; Milošević, M.M.; Antov, M.G. Protein Nanoparticles From Chickpea Albumin, Globulin, and Glutelin—The Impact of Heat Treatment on Secondary Structure, Surface Hydrophobicity, and Properties. Food Bioprocess Technol. 2025, 18, 535–547. [Google Scholar] [CrossRef] [Scilit]
  11. Grasso, N.; Lynch, N.L.; Arendt, E.K.; O’Mahony, J.A. Chickpea Protein Ingredients: A Review of Composition, Functionality, and Applications. Compr. Rev. Food Sci. Food Saf. 2022, 21, 435–452. [Google Scholar] [CrossRef] [Scilit]
  12. Mahdavian Mehr, H.; Koocheki, A. Effect of Atmospheric Cold Plasma on Structure, Interfacial and Emulsifying Properties of Grass Pea (Lathyrus sativus L.) Protein Isolate. Food Hydrocoll. 2020, 106, 105899. [Google Scholar] [CrossRef] [Scilit]
  13. Chang, L.; Lan, Y.; Bandillo, N.; Ohm, J.-B.; Chen, B.; Rao, J. Plant Proteins from Green Pea and Chickpea: Extraction, Fractionation, Structural Characterization and Functional Properties. Food Hydrocoll. 2022, 123, 107165. [Google Scholar] [CrossRef] [Scilit]
  14. Tang, C.-H. Emulsifying Properties of Soy Proteins: A Critical Review with Emphasis on the Role of Conformational Flexibility. Crit. Rev. Food Sci. Nutr. 2017, 57, 2636–2679. [Google Scholar] [CrossRef] [Scilit]
  15. Guo, J.; Yang, X.-Q.; He, X.-T.; Wu, N.-N.; Wang, J.-M.; Gu, W.; Zhang, Y.-Y. Limited Aggregation Behavior of β-Conglycinin and Its Terminating Effect on Glycinin Aggregation during Heating at pH 7.0. J. Agric. Food Chem. 2012, 60, 3782–3791. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  16. Tang, Y.R.; Ghosh, S. Canola Protein Thermal Denaturation Improved Emulsion-Templated Oleogelation and Its Cake-Baking Application. RSC Adv. 2021, 11, 25141–25157. [Google Scholar] [CrossRef] [Scilit]
  17. Tavernier, I.; Patel, A.R.; Van Der Meeren, P.; Dewettinck, K. Emulsion-Templated Liquid Oil Structuring with Soy Protein and Soy Protein: κ-Carrageenan Complexes. Food Hydrocoll. 2017, 65, 107–120. [Google Scholar] [CrossRef] [Scilit]
  18. Patel, A.R.; Rajarethinem, P.S.; Cludts, N.; Lewille, B.; De Vos, W.H.; Lesaffer, A.; Dewettinck, K. Biopolymer-Based Structuring of Liquid Oil into Soft Solids and Oleogels Using Water-Continuous Emulsions as Templates. Langmuir 2015, 31, 2065–2073. [Google Scholar] [CrossRef] [Scilit]
  19. Keshanidokht, S.; Kumar, S.; Thulstrup, P.W.; Via, M.A.; Clausen, M.P.; Risbo, J. Thermo-Responsive Behavior of Glycerol-Plasticized Oleogels Stabilized by Zein. Food Hydrocoll. 2023, 139, 108582. [Google Scholar] [CrossRef] [Scilit]
  20. Feichtinger, A.; Nibbelink, D.G.; Poppe, S.; Bozzo, L.; Landman, J.; Scholten, E. Protein Oleogels Prepared by Solvent Transfer Method with Varying Protein Sources. Food Hydrocoll. 2022, 132, 107821. [Google Scholar] [CrossRef] [Scilit]
  21. Mohanan, A.; Tang, Y.R.; Nickerson, M.T.; Ghosh, S. Oleogelation Using Pulse Protein-Stabilized Foams and Their Potential as a Baking Ingredient. RSC Adv. 2020, 10, 14892–14905. [Google Scholar] [CrossRef] [Scilit]
  22. Chen, H.; Wang, Q.; Liu, S.; Rao, Z.; Lei, X.; Zhao, J.; Lei, L.; Zeng, K.; Ming, J. The Characterization of Zein/Xanthan Gum Composite Oleogel: A Comparative Study of Bioactive Substances in Interface and Oil Phases. Food Hydrocoll. 2025, 159, 110613. [Google Scholar] [CrossRef] [Scilit]
  23. Meng, Z.; Qi, K.; Guo, Y.; Wang, Y.; Liu, Y. Physical Properties, Microstructure, Intermolecular Forces, and Oxidation Stability of Soybean Oil Oleogels Structured by Different Cellulose Ethers. Eur. J. Lipid Sci. Technol. 2018, 120, 1700287. [Google Scholar] [CrossRef] [Scilit]
  24. Perović, M.N.; Pajin, B.S.; Antov, M.G. The Effect of Enzymatic Pretreatment of Chickpea on Functional Properties and Antioxidant Activity of Alkaline Protein Isolate. Food Chem. 2022, 374, 131809. [Google Scholar] [CrossRef] [Scilit]
  25. Chang, Y.-W.; Alli, I.; Konishi, Y.; Ziomek, E. Characterization of Protein Fractions from Chickpea (Cicer arietinum L.) and Oat (Avena sativa L.) Seeds Using Proteomic Techniques. Food Res. Int. 2011, 44, 3094–3104. [Google Scholar] [CrossRef] [Scilit]
  26. Bradford, M.M. A Rapid and Sensitive Method for the Quantitation of Microgram Quantities of Protein Utilizing the Principle of Protein-Dye Binding. Anal. Biochem. 1976, 72, 248–254. [Google Scholar] [CrossRef]
  27. Mohamed, A.; Hojilla-Evangelista, M.P.; Peterson, S.C.; Biresaw, G. Barley Protein Isolate: Thermal, Functional, Rheological, and Surface Properties. J. Am. Oil Chem. Soc. 2007, 84, 281–288. [Google Scholar] [CrossRef] [Scilit]
  28. Ke, L.; Xu, Y.; Gao, G.; Wang, H.; Yu, Z.; Zhou, J.; Rao, P.; Wang, Q.; Yu, J. Catalase to Demulsify Oil-in-Water Fish Oil-Polysorbate Emulsion and Affect Lipid Oxidation. Food Res. Int. 2020, 133, 109169. [Google Scholar] [CrossRef] [Scilit]
  29. Miao, W.; McClements, D.J.; Zhang, Z.; Lin, Q.; Ji, H.; Wang, J.; Jin, Z.; Li, G.; Jiang, L.; Wen, J.; et al. Fabrication and Characterization of Emulsion-Template Oleogels Assembled from Octenyl Succinic Anhydride Starch/Chitosan Electrostatic Complexes. Food Hydrocoll. 2024, 151, 109882. [Google Scholar] [CrossRef] [Scilit]
  30. Wang, Y.; Wang, Y.; Li, K.; Bai, Y.; Li, B.; Xu, W. Effect of High Intensity Ultrasound on Physicochemical, Interfacial and Gel Properties of Chickpea Protein Isolate. LWT 2020, 129, 109563. [Google Scholar] [CrossRef] [Scilit]
  31. Li, H.; Wu, X.; Lin, L.; Zhou, X.; Wu, W. Effect of Epigallocatechin-3-Gallate Modification on the Structure, Hydrophilicity and Pickering Emulsion Stability of Rice Bran Protein-Polysaccharide-Phenol Natural Complex. Food Chem. 2025, 482, 144175. [Google Scholar] [CrossRef] [Scilit]
  32. Bourouis, I.; Sun, M.; Li, H.; Pang, Z.; Liu, X. Formation and Characterization of Soy Protein Isolate-Chitosan-Epigallocatechin Gallate Ternary Complexes: Molecular Interactions and Functional Properties. Carbohydr. Polym. 2026, 371, 124474. [Google Scholar] [CrossRef] [Scilit]
  33. Viseu, M.I.; Carvalho, T.I.; Costa, S.M.B. Conformational Transitions in B-Lactoglobulin Induced by Cationic Amphiphiles: Equilibrium Studies. Biophys. J. 2004, 86, 2392–2402. [Google Scholar] [CrossRef] [Scilit]
  34. Lakemond, C.M.M.; De Jongh, H.H.J.; Hessing, M.; Gruppen, H.; Voragen, A.G.J. Soy Glycinin: Influence of pH and Ionic Strength on Solubility and Molecular Structure at Ambient Temperatures. J. Agric. Food Chem. 2000, 48, 1985–1990. [Google Scholar] [CrossRef] [Scilit]
  35. Adiamo, O.Q.; Netzel, M.E.; Hoffman, L.C.; Gidley, M.J.; Osborne, S.; Sultanbawa, Y. Structure—Function Relationship of Australian Acacia Seed Protein Concentrates: Amino Acid Composition, in Vitro Protein Digestibility and Molecular Properties. Food Biosci. 2023, 51, 102339. [Google Scholar] [CrossRef] [Scilit]
  36. Niu, F.; Dong, Y.; Shen, F.; Wang, J.; Liu, Y.; Su, Y.; Xu, R.; Wang, J.; Yang, Y. Phase Separation Behavior and Structural Analysis of Ovalbumin–Gum Arabic Complex Coacervation. Food Hydrocoll. 2015, 43, 1–7. [Google Scholar] [CrossRef] [Scilit]
  37. Pan, Y.; Liu, S.; Han, Z.; Li, H.; Lai, H.; Shao, J.-H.; Xiao, N.; Xia, W. The Beeswax-Based Fish Oil Oleogels Adjusting the Stabilization Mechanism and Lipid Digestion of Bi-Layer Emulsions: Spatial Conformation and Molecular Interaction. Food Hydrocoll. 2025, 158, 110561. [Google Scholar] [CrossRef] [Scilit]
  38. Niu, H.; Chen, X.; Fu, X.; Zhang, B.; Dou, Z.; Huang, Q. Pectin-Stabilized Emulsions: Structure-Emulsification Relationships, Covalent and Non-Covalent Modifications, and Future Trends. Trends Food Sci. Technol. 2025, 159, 104986. [Google Scholar] [CrossRef] [Scilit]
  39. Zhu, J.; Liu, L.; Li, X.; Zhang, Q.; Wang, Z.; Chen, N.; Wang, H.; Xie, F.; Qi, B.; Jiang, L. Construction of Soybean Oil Bodies–Xanthan Gum Composite Oleogels by Emulsion-Templated Method: Preparation, Characterization, and Stability Analysis. Food Hydrocoll. 2024, 149, 109526. [Google Scholar] [CrossRef] [Scilit]
  40. Wu, Y.; You, F.; Zheng, Y. Inhibited Hydration of Xanthan Gum under Synergistic High-pH/Ca2+ Conditions: Rheological Behavior and Molecular Dynamics Simulations. Carbohydr. Polym. 2026, 374, 124704. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  41. Zhao, T.; Huang, K.; Luo, Y.; Li, Y.; Cheng, N.; Mei, X. Preparation and Characterization of High Internal Phase Pickering Emulsions Stabilized by Hordein-Chitosan Composite Nanoparticles. Colloids Surf. A Physicochem. Eng. Asp. 2023, 659, 130766. [Google Scholar] [CrossRef] [Scilit]
  42. Ebrahimi, R.; Fathi, M.; Ghoddusi, H.B. Pickering Emulsions Stabilized by Cellulose Nanocrystals Extracted from Hazelnut Shells: Production and Stability under Different Harsh Conditions. Int. J. Biol. Macromol. 2024, 258, 128982. [Google Scholar] [CrossRef] [Scilit]
  43. Hettiarachchy, N.S.; Ziegler, G.R. Protein Functionality in Food Systems; CRC Press: Boca Raton, FL, USA, 1994; ISBN 978-1-4822-9351-7. [Google Scholar]
  44. Turgeon, S.L.; Beaulieu, M.; Schmitt, C.; Sanchez, C. Protein–Polysaccharide Interactions: Phase-Ordering Kinetics, Thermodynamic and Structural Aspects. Curr. Opin. Colloid Interface Sci. 2003, 8, 401–414. [Google Scholar] [CrossRef] [Scilit]
  45. Doublier, J.-L.; Garnier, C.; Renard, D.; Sanchez, C. Protein–Polysaccharide Interactions. Curr. Opin. Colloid Interface Sci. 2000, 5, 202–214. [Google Scholar] [CrossRef] [Scilit]
  46. Stoll, S.; Chodanowski, P. Polyelectrolyte Adsorption on an Oppositely Charged Spherical Particle. Chain Rigidity Effects. Macromolecules 2002, 35, 9556–9562. [Google Scholar] [CrossRef] [Scilit]
  47. Akinchina, A.; Linse, P. Monte Carlo Simulations of Polyion−Macroion Complexes. 1. Equal Absolute Polyion and Macroion Charges. Macromolecules 2002, 35, 5183–5193. [Google Scholar] [CrossRef] [Scilit]
  48. Ma, X.; Habibi, M.; Sagis, L.M.C. Air-Water Interfacial and Foaming Properties of Lupin Protein-Polysaccharide Soluble Complexes: Role of Physicochemical Properties, Morphological Characteristics, and Flexibility. Food Hydrocoll. 2025, 165, 111247. [Google Scholar] [CrossRef] [Scilit]
  49. Jiang, L.; Wang, Q.; Rao, Z.; Lei, X.; Zhao, J.; Lei, L.; Ming, J. Formulation and Characterization of Bigels Utilizing Whey Protein and Polysaccharides: Potential Applications as Cream Analogues. Food Hydrocoll. 2024, 152, 109884. [Google Scholar] [CrossRef] [Scilit]
  50. Li, F.; Li, X.; Huang, K.; Luo, Y.; Mei, X. Preparation and Characterization of Pickering Emulsion Stabilized by Hordein-Chitosan Complex Particles. J. Food Eng. 2021, 292, 110275. [Google Scholar] [CrossRef] [Scilit]
  51. Shan, G.; Cui, M.; Wang, X.; Liang, X.; Xu, Z.; Zhang, Y.; Sui, X. Fabrication of Enhanced Aerogel Template Oleogels with Enzyme-Hydrolyzed Soy Protein Isolate and Covalent Cross-Linking. Int. J. Biol. Macromol. 2024, 275, 133674. [Google Scholar] [CrossRef] [Scilit]
  52. Lunkad, R.; Barroso Da Silva, F.L.; Košovan, P. Both Charge-Regulation and Charge-Patch Distribution Can Drive Adsorption on the Wrong Side of the Isoelectric Point. J. Am. Chem. Soc. 2022, 144, 1813–1825. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  53. Damodaran, S. Protein Stabilization of Emulsions and Foams. J. Food Sci. 2005, 70, R54–R66. [Google Scholar] [CrossRef] [Scilit]
  54. Ma, X.; Hou, F.; Zhao, H.; Wang, D.; Chen, W.; Miao, S.; Liu, D. Conjugation of Soy Protein Isolate (SPI) with Pectin by Ultrasound Treatment. Food Hydrocoll. 2020, 108, 106056. [Google Scholar] [CrossRef] [Scilit]
  55. Tirgarian, B.; Farmani, J.; Farahmandfar, R.; Milani, J.M.; Van Bockstaele, F. Colloidal Network Oleogels Structured by Sonothermal Conjugates of Sodium Caseinate and Anionic Gums. Food Hydrocoll. 2023, 140, 108624. [Google Scholar] [CrossRef] [Scilit]
  56. Huang, Z.; Li, T.; Zhao, J.; Zhang, H.; Fan, M.; Li, Y.; Qian, H.; Wang, L. Characteristics of Emulsions Stabilized by Mung Bean Protein Fractions: Protein Aggregate Behaviours and Interface Properties. Food Hydrocoll. 2026, 171, 111865. [Google Scholar] [CrossRef] [Scilit]
  57. Skoglund, T. Theoretical Analysis of Methods to Measure Homogenisation Efficiency, Based on Centrifugation and Droplet Size Distribution. Int. Dairy J. 2025, 169, 106309. [Google Scholar] [CrossRef] [Scilit]
  58. Alade, O.S.; Mahmoud, M.; Al Shehri, D.A.; Sultan, A.S. Rapid Determination of Emulsion Stability Using Turbidity Measurement Incorporating Artificial Neural Network (ANN): Experimental Validation Using Video/Optical Microscopy and Kinetic Modeling. ACS Omega 2021, 6, 5910–5920. [Google Scholar] [CrossRef] [Scilit]
  59. Li, H.; Li, F.; Wu, X.; Wu, W. Effect of Rice Bran Rancidity on the Emulsion Stability of Rice Bran Protein and Structural Characteristics of Interface Protein. Food Hydrocoll. 2021, 121, 107006. [Google Scholar] [CrossRef] [Scilit]
  60. Liu, X.; Liu, Y.; Meng, F.; Wan, Z.; Yang, X. Oleogels by Zein-Tea Saponin Complex Nanoparticles-Stabilized Pickering Emulsions Templating and Application as Animal Fat Replacers in Beef Burger. LWT 2024, 203, 116328. [Google Scholar] [CrossRef] [Scilit]
  61. Xu, Y.; Sun, H.; Lv, J.; Wang, Y.; Zhang, Y.; Wang, F. Effects of Polysaccharide Thickening Agent on the Preparation of Walnut Oil Oleogels Based on Methylcellulose: Characterization and Delivery of Curcumin. Int. J. Biol. Macromol. 2023, 232, 123291. [Google Scholar] [CrossRef] [Scilit]
  62. Barroso, N.G.; Okuro, P.K.; Ribeiro, A.P.B.; Cunha, R.L. Tailoring Properties of Mixed-Component Oleogels: Wax and Monoglyceride Interactions Towards Flaxseed Oil Structuring. Gels 2020, 6, 5. [Google Scholar] [CrossRef] [Scilit]
  63. Jiang, H.; Qi, J. Structuring Oleogels with Citrus Fiber as the Sole Gelling Agent: Precise Modulation of the Network Structure via Enzymatic Method. Food Hydrocoll. 2025, 174, 112309. [Google Scholar] [CrossRef] [Scilit]
  64. Sun, Y.; Zhong, M.; Zhao, X.; Li, Y.; Qi, B.; Jiang, L. Stability and Digestion Characteristics of Pickering High Internal Phase Emulsions Formed by Acid-Induced Soy Lipophilic Protein, β-Conglycinin, and Globulin. LWT 2022, 153, 112554. [Google Scholar] [CrossRef] [Scilit]
  65. Jin, Y.; Li, X.; Wu, J.; Xiao, J.; Mei, X. Oleogels Based on Protein Fibrils-Hordein-Xanthan Gum Complexes: Formation, Structure, and Physicochemical Properties. Food Res. Int. 2025, 208, 116244. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  66. Brito, G.B.; Peixoto, V.O.D.S.; Martins, M.T.; Rosário, D.K.A.; Ract, J.N.; Conte-Júnior, C.A.; Torres, A.G.; Castelo-Branco, V.N. Development of Chitosan-Based Oleogels via Crosslinking with Vanillin Using an Emulsion Templated Approach: Structural Characterization and Their Application as Fat-Replacer. Food Struct. 2022, 32, 100264. [Google Scholar] [CrossRef] [Scilit]
  67. Wang, Q.; Rao, Z.; Chen, Y.; Jiang, L.; Lei, X.; Zhao, J.; Li, F.; Lei, L.; Ming, J. Fabrication and Characterization of Oleogels Stabilized by Metal-Phenolic Network Coatings-Decorated Zein Nanoparticles. Food Chem. 2024, 430, 137025. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  68. Yang, S.; Jin, Y.; Li, F.; Shi, J.; Liang, J.; Mei, X. Pickering Emulsion Stabilized by Hordein–Whey Protein Isolate Complex: Delivery System of Quercetin. Foods 2024, 13, 665. [Google Scholar] [CrossRef] [Scilit]
  69. Huang, Y.; Ma, T.; Li, C.; McClements, D.J. Plant-Based Emulsion Gels Assembled from Potato Protein, β-Glucan and Olive Oil: Structure, Functionality, and Application as Low-Fat Butter Replacers. Food Hydrocoll. 2026, 174, 112439. [Google Scholar] [CrossRef] [Scilit]
  70. Meng, Z.; Qi, K.; Guo, Y.; Wang, Y.; Liu, Y. Macro-Micro Structure Characterization and Molecular Properties of Emulsion-Templated Polysaccharide Oleogels. Food Hydrocoll. 2018, 77, 17–29. [Google Scholar] [CrossRef] [Scilit]
  71. Liu, N.; Lin, P.; Zhang, K.; Li, D.; Yang, D.; Katsuyoshi, N.; Zhao, M.; Yao, X. Konjac Glucomannan Promoted Fabrication of Diacylglycerol Oil-Based Oleogels through Emulsion-Templated Approach: Comparison with Triacylglycerol Oleogels. Int. J. Biol. Macromol. 2023, 251, 126420. [Google Scholar] [CrossRef] [Scilit]
  72. Patel, A.R.; Dewettinck, K. Comparative Evaluation of Structured Oil Systems: Shellac Oleogel, HPMC Oleogel, and HIPE Gel. Eur. J. Lipid Sci. Technol. 2015, 117, 1772–1781. [Google Scholar] [CrossRef] [Scilit]
  73. Abdolmaleki, K.; Alizadeh, L.; Nayebzadeh, K.; Hosseini, S.M.; Shahin, R. Oleogel Production Based on Binary and Ternary Mixtures of Sodium Caseinate, Xanthan Gum, and Guar Gum: Optimization of Hydrocolloids Concentration and Drying Method. J. Texture Stud. 2020, 51, 290–299. [Google Scholar] [CrossRef] [Scilit] [PubMed]
  74. Pang, M.; Wang, X.; Cao, L.; Shi, Z.; Lei, Z.; Jiang, S. Structure and Thermal Properties of Β-sitosterol-beeswax-sunflower Oleogels. Int. J. Food Sci. Technol. 2020, 55, 1900–1908. [Google Scholar] [CrossRef] [Scilit]
  75. Naderi, M.; Farmani, J.; Rashidi, L. Structuring of Chicken Fat by Monoacylglycerols. J. Am. Oil Chem. Soc. 2016, 93, 1221–1231. [Google Scholar] [CrossRef] [Scilit]
Foods 15 00905 g001
Figure 1. SDS-PAGE of CPP, ALB, GLO, and GLU (A). Surface hydrophobicity of proteins and their corresponding complexes (B). Different letters represent a significant difference (p < 0.05). Different lowercase and capital letters respectively indicate significant differences among the proteins and the complexes (p < 0.05). “*” indicates a significant difference between the proteins and their corresponding complexes (* p < 0.05, ** p < 0.01).
Figure 1. SDS-PAGE of CPP, ALB, GLO, and GLU (A). Surface hydrophobicity of proteins and their corresponding complexes (B). Different letters represent a significant difference (p < 0.05). Different lowercase and capital letters respectively indicate significant differences among the proteins and the complexes (p < 0.05). “*” indicates a significant difference between the proteins and their corresponding complexes (* p < 0.05, ** p < 0.01).
Foods 15 00905 g001
Foods 15 00905 g002
Figure 2. Fluorescence spectra of the four proteins, protein–XG complexes, and XG.
Figure 2. Fluorescence spectra of the four proteins, protein–XG complexes, and XG.
Foods 15 00905 g002
Foods 15 00905 g003
Figure 3. Particle size and ζ-potential of the four proteins, protein–XG complexes and XG (A,B). FTIR spectra of CPP, XG, and their complexes (C). Different lowercase and capital letters respectively indicate significant differences among the proteins and the complexes (p < 0.05). “*” indicates a significant difference between the proteins and their corresponding complexes (* p < 0.05).
Figure 3. Particle size and ζ-potential of the four proteins, protein–XG complexes and XG (A,B). FTIR spectra of CPP, XG, and their complexes (C). Different lowercase and capital letters respectively indicate significant differences among the proteins and the complexes (p < 0.05). “*” indicates a significant difference between the proteins and their corresponding complexes (* p < 0.05).
Foods 15 00905 g003
Foods 15 00905 g004
Figure 4. Appearance of the emulsions (A). D [4,3] and centrifugal stability of the emulsions (B). Different letters indicate a significant difference (p < 0.05).
Figure 4. Appearance of the emulsions (A). D [4,3] and centrifugal stability of the emulsions (B). Different letters indicate a significant difference (p < 0.05).
Foods 15 00905 g004
Foods 15 00905 g005
Figure 5. Fluorescence microstructure of the emulsions.
Figure 5. Fluorescence microstructure of the emulsions.
Foods 15 00905 g005
Foods 15 00905 g006
Figure 6. Appearance of the oleogels (A). OBC of the oleogels (B). Different letters represent a significant difference (p < 0.05).
Figure 6. Appearance of the oleogels (A). OBC of the oleogels (B). Different letters represent a significant difference (p < 0.05).
Foods 15 00905 g006
Foods 15 00905 g007
Figure 7. CLSM images of the oleogels.
Figure 7. CLSM images of the oleogels.
Foods 15 00905 g007
Foods 15 00905 g008
Figure 8. Formation mechanism of the oleogel network.
Figure 8. Formation mechanism of the oleogel network.
Foods 15 00905 g008
Foods 15 00905 g009
Figure 9. Amplitude sweeps for oleogels (A). Frequency sweeps for oleogels (B). Apparent viscosity (C). Thixotropic test curves (D). Temperature sweep curves (E). DSC spectra of oleogels during the heating process (F).
Figure 9. Amplitude sweeps for oleogels (A). Frequency sweeps for oleogels (B). Apparent viscosity (C). Thixotropic test curves (D). Temperature sweep curves (E). DSC spectra of oleogels during the heating process (F).
Foods 15 00905 g009
Table 1. Percent thixotropic recovery of oleogels.
Table 1. Percent thixotropic recovery of oleogels.
Thixotropic Recovery (%)
CPP-XO71.91 ± 0.95 b
ALB-XO69.35 ± 1.31 c
GLO-XO65.51 ± 2.94 d
GLU-XO75.45 ± 1.84 a
Different letters indicate a significant difference (p < 0.05).
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Li, X.; Yang, S.; Wu, J.; Jin, Y.; Mei, X. Oleogels Based on Chickpea Protein Fractions–Xanthan Gum Complexes: Preparation and Characterization. Foods 2026, 15, 905. https://doi.org/10.3390/foods15050905

AMA Style

Li X, Yang S, Wu J, Jin Y, Mei X. Oleogels Based on Chickpea Protein Fractions–Xanthan Gum Complexes: Preparation and Characterization. Foods. 2026; 15(5):905. https://doi.org/10.3390/foods15050905

Chicago/Turabian Style

Li, Xiaomeng, Songqi Yang, Jingwen Wu, Yunan Jin, and Xiaohong Mei. 2026. "Oleogels Based on Chickpea Protein Fractions–Xanthan Gum Complexes: Preparation and Characterization" Foods 15, no. 5: 905. https://doi.org/10.3390/foods15050905

APA Style

Li, X., Yang, S., Wu, J., Jin, Y., & Mei, X. (2026). Oleogels Based on Chickpea Protein Fractions–Xanthan Gum Complexes: Preparation and Characterization. Foods, 15(5), 905. https://doi.org/10.3390/foods15050905

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop