Effect of Wall-Material Assembly Sequence on Ovalbumin–Chitosan Nanoparticles for Antarctic Krill Peptide Delivery

TL;DR

This study shows that changing the order of materials in nanoparticles affects their size, stability, and ability to deliver Antarctic krill peptides for health benefits.

Contribution

The novelty lies in demonstrating how wall-material assembly sequence impacts nanoparticle properties and bioactive peptide delivery efficiency.

Findings

CS/OVA-AKP nanoparticles had better physical properties and higher encapsulation efficiency than OVA/CS-AKP.

CS/OVA-AKP showed greater stability and protection of peptides during simulated digestion.

CS/OVA-AKP exhibited higher hypoglycemic activity by inhibiting key digestive enzymes.

Abstract

The objective of this study was to explore the effect of the assembly sequences of wall materials on the structure and properties of Antarctic krill peptide (AKP)-loaded ovalbumin (OVA)–chitosan (CS) nanoparticles (NPs). Two AKP-loaded NPs (CS/OVA-AKP and OVA/CS-AKP) were prepared by changing the sequences of OVA and CS. The results confirmed that CS/OVA-AKP had a smaller particle size (291 nm vs. 320 nm), lower polydispersity index (0.233 vs. 0.282), higher absolute zeta potential (34.4 mV vs. 32.1 mV), and higher encapsulation efficiency (81.6% vs. 75.4%) than OVA/CS-AKP. X-ray diffraction analysis confirmed that AKP was encapsulated in an amorphous state within the NPs. Fourier transform infrared spectroscopy and three-dimensional (3D) fluorescence spectroscopy revealed that electrostatic interactions, hydrogen bonding, and hydrophobic interactions were the primary driving forces for nanoparticle formation, with CS/OVA-AKP demonstrating a stronger OVA fluorescence quenching effect. Compared with OVA/CS-AKP, CS/OVA-AKP exhibited better redispersibility, and CS/OVA-AKP showed greater stability under various environmental factors (thermal treatment, salt concentration, pH, and storage time). During simulated gastrointestinal digestion, CS/OVA-AKP effectively protected AKP from gastric degradation and showed a higher AKP release rate in simulated intestinal fluid (61.1%) than OVA/CS-AKP (53.0%). The release followed the Korsmeyer–Peppas model, with OVA/CS-AKP exhibiting non-Fickian diffusion (n = 0.7500), and CS/OVA-AKP approached Case II transport (n = 0.9889), indicating erosion-controlled release behavior. CS/OVA-AKP also demonstrated higher hypoglycemic activity, with inhibition rates of 41.1%, 37.5%, and 36.1% for α-glucosidase, α-amylase, and DPP-IV, respectively. These findings underscore the important influence of wall-material assembly sequences on the structure and properties of AKP-loaded NPs, offering valuable insights for the development of bioactive peptide delivery systems.

Figures

Full text

1. Introduction

Antarctic krill peptide (AKP) is a bioactive peptide derived from marine sources that exhibits a variety of physiological regulatory activities, involving antioxidant, hypoglycemic, and immunomodulatory effects, indicating its considerable potential in addressing chronic metabolic diseases, including diabetes [1,2]. Diabetes mellitus is a multifaceted metabolic condition marked by elevated blood glucose levels, potentially resulting in a range of complications. It is one of the fastest-growing global public health issues [3]. According to data from the International Diabetes Federation (IDF), the global number of individuals living with diabetes reached 537 million in 2021 and continues to rise [4]. Previous studies have demonstrated that AKP possesses strong hypoglycemic efficacy and the ability to enhance insulin sensitivity [5,6]. Nevertheless, the practical application of AKP is constrained by its inadequate stability during food processing, long-term storage, and gastrointestinal digestion. Fluctuations in pH, thermal treatment, and exposure to the harsh digestive environment can induce peptide chain cleavage and conformational changes, leading to a marked reduction in its bioactivity [7,8]. Therefore, it is of significant importance to develop biocompatible carrier systems that improve the stability of AKP and preserve its biological activity.

The formation of nanoparticles by interactions between proteins and polysaccharides is an efficient method for developing biocompatible and biosafe delivery systems [9,10]. Ovalbumin (OVA), an animal-derived amphipathic globular protein, is widely used as a delivery carrier due to its excellent nutritional value [11]. However, OVA as a single delivery vehicle has several drawbacks, including poor thermal stability, sensitivity to salt ions, a tendency to aggregate near its isoelectric point (pI = 4.5), and susceptibility to proteolytic hydrolysis. These issues can lead to leakage or degradation of encapsulated bioactive substances, significantly limiting their effectiveness [12,13]. Additionally, OVA is a well-known food-derived allergenic protein that can induce immune responses or allergic reactions in susceptible individuals [14]. Common processing methods such as physical treatment, chemical modification, and biological treatment can effectively reduce the allergenicity of OVA [15]. Bioactive polysaccharides exhibit excellent anti-allergic activity. They provide a more valuable application direction for addressing the allergenicity issue of OVA [16]. To address these limitations of OVA, the incorporation of polysaccharides has been shown to significantly improve nanoparticles’ stability and functional characteristics. Compared to proteins, polysaccharides demonstrate good stability across a broad pH range and effectively protect active substances from digestion in gastric fluids [17]. Chitosan (CS), as the only natural cationic polysaccharide, not only exhibits good biocompatibility and adhesiveness, but the dense hydrogen bond network formed between its molecular chains also provides steric hindrance, reducing particle aggregation [18,19]. Studies have shown that CS could form a compact core-shell structure with negatively charged protein through electrostatic interactions, which helped inhibit proteolytic degradation of protein [20,21]. In simulated digestive environments, CS-coated nanoparticles significantly delayed pepsin-induced degradation of the core protein and improved the intestinal release rate of bioactive peptides [22]. Additionally, the cationic properties of CS enhanced nanoparticle adhesion to intestinal mucosa, thereby prolonging the retention time of bioactive peptides at the target site [23,24].

The method used to prepare nanoparticles, along with the characteristics of the wall material, plays a vital part in defining the structure and properties of the nanoparticles, thereby regulating the delivery efficiency of bioactive peptides. Research indicates that the wall material’s blending sequences may affect nanoparticle assembly behavior by altering the three-dimensional conformation of proteins and intermolecular interactions, including hydrophobic associations, hydrogen bonds, and electrostatic attractions [25,26]. The preparation of nanoparticles is a key step in the engineering of delivery systems. Optimizing the assembly sequences can improve both the drug loading efficiency and the stability of the delivery system by modulating the interactions between the wall and core materials. Chen et al. [27] showed that modifying the blending sequences of the wall material in the antisolvent precipitation method could lead to nanoparticles exhibiting better storage stability and in vitro bioavailability. Yang et al. [28] found that different blending sequences affected the distribution of lecithin molecules on the nanoparticle surface, which in turn affected the curcumin encapsulation efficiency. However, although previous studies have shown that the assembly sequences of wall materials affect the encapsulation and release of small hydrophobic nutraceuticals such as curcumin, it remained unclear whether such sequence-dependent assembly mechanisms similarly regulate the protection efficiency, functional properties, and release kinetics of bioactive peptides such as AKP. Moreover, studies on OVA-CS composite nanoparticles for AKP delivery are limited, particularly with respect to the effects of wall-material assembly sequences on their structural and functional characteristics. Therefore, this study investigated the encapsulation of AKP by OVA and CS, exploring how the assembly sequences affect the structure and properties of nanoparticles. The findings are intended to serve as a foundation for developing delivery systems based on protein-polysaccharide nanoparticles for active peptides.

2. Materials and Methods

2.1. Materials

Defatted Antarctic krill powder was obtained from Qingdao Kangjing Marine Biotechnology Co., Ltd. (Qingdao, China). Ovalbumin (Mw: 44 kDa), chitosan (deacetylation degree: 90%, Mw: 100 kDa), pepsin, and pancreatin were sourced from Shanghai McLean Biochemical Technology Co., Ltd. (Shanghai, China). The composite protease was purchased from Tianjin Nuoao Biotechnology Co., Ltd. (Tianjin, China). The DPP-IV inhibitor screening assay kit was supplied by Abon Antibody (Shanghai, China) Trading Co., Ltd. (Shanghai, China). The remaining reagents utilised were all of analytical grade.

2.2. Preparation of Antarctic Krill Peptides (AKP)

A phosphate buffer solution (pH 7, 0.2 mol/L) was used to disperse defatted Antarctic krill powder at a 1:6 (m/v) ratio. The blend was thoroughly stirred (300× g, 10 min) and subsequently sonicated (360 W, 10 min). A composite protease, equivalent to 7% of the de-fatted Antarctic krill powder mass (with an enzyme activity of 6.22 × 10^4^ U/g), was incorporated, the system’s pH was set to 7, and enzymatic hydrolysis was carried out in a water bath (50 °C, 6 h). Subsequently, the enzyme was deactivated by raising the temperature (90 °C, 15 min). After centrifuging (10,000× g, 15 min), the supernatant was subjected to ultrafiltration membranes to obtain the Antarctic krill peptide solution with a molecular weight under 3 kDa. The solution underwent freeze-drying (48 h) to yield the AKP [29].

2.3. Amino Acid Analysis and Molecular Weight (Mw) Distribution

The detailed procedures for amino acid analysis and Mw distribution determination of AKP were provided in the Supplementary Materials.

2.4. Preparation of AKP-Loaded Nanoparticles

The AKP-loaded nanoparticles (NPs) were prepared utilizing OVA and CS as wall materials by electrostatic self-assembly [30]. OVA and CS were solubilized in deionized water and acetic acid solution (1%, v/v), respectively, to prepare OVA (1 mg/mL) and CS (1 mg/mL) solutions. The mixtures were stirred overnight to ensure full hydration. AKP was dissolved in deionized water and mixed using magnetic stirring (400× g, 30 min) to achieve a 1 mg/mL solution. To make the OVA-AKP suspension, the AKP solution was gradually introduced into the OVA solution at a 1:1 (v/v) ratio and agitated at room temperature (400× g, 2 h). The pH was adjusted to 5.5 after adding this suspension dropwise to the CS solution. By stirring the mixture at room temperature (400× g, 2 h), the CS/OVA-AKP suspension was prepared. The mass ratio of CS to OVA was set at 1:3, 1:2, 1:1, 2:1, and 3:1, respectively. The resulting nanoparticle suspensions were named CS/OVA-AKP 1:3, CS/OVA-AKP 1:2, CS/OVA-AKP 1:1, CS/OVA-AKP 2:1, and CS/OVA-AKP 3:1. The NPs were obtained by freeze-drying a portion of the suspension for 48 h.

The appropriate ratios were chosen by considering particle size, polydispersity index (PDI), zeta potential, encapsulation efficiency (EE), and loading efficiency (LC). Under the same preparation method and experimental conditions, OVA/CS-AKP suspensions and NPs were prepared by varying the order of addition of OVA and CS.

2.5. Particle Size, PDI, and Zeta Potential

Dynamic light scattering (DLS) was employed to assess the particle size, PDI, and zeta potential of AKP-loaded nanoparticle suspensions, with a Malvern particle size analyzer (Nano-ZS 90, Malvern Instruments Ltd., Malvern, UK) employed for the analysis [31].

2.6. Encapsulation Efficiency (EE) and Loading Capacity (LC)

The EE and LC were measured using Han et al.’s method [32]. Briefly, the nanoparticle suspensions were placed into ultrafiltration centrifuge tubes (molecular weight cutoff: 3 kDa) and centrifuged (4 °C, 4500× g, 30 min). The concentration of AKP in the filtrate was quantified utilizing an ultraviolet spectrophotometer (UV-1800, Shimadzu Corporation, Kyoto, Japan) at 265 nm, based on a standard curve of AKP (R^2^ ≥ 0.999). The EE and LC were computed utilizing the subsequent equations:

$$[eqn]$$ $$[eqn]$$

2.7. Structural Characterization

2.7.1. Fourier Transform Infrared Spectroscopy (FTIR)

The FTIR analysis was conducted using a Nicolet iS10 (Thermo Scientific, Waltham, MA, USA) according to Li et al. [33]. Samples were scanned 32 times at 4 cm^−1^ resolution over 400–4000 cm^−1^ wavelength.

2.7.2. Three-Dimensional (3D) Fluorescence Spectroscopy

The three-dimensional (3D) fluorescence spectroscopy was acquired according to Cheng et al. [34], with samples diluted to 0.01 mg/mL OVA. Excitation and emission wavelengths were 200–350 and 200–500 nm, excitation and emission intervals were 5 nm, voltage was 700 V, and scanning speed was 12,000 nm/min.

2.7.3. Scanning Electron Microscopy (SEM)

The freeze-dried samples’ microscopic morphology was examined by scanning electron microscopy (SEM, SU-5000, Hitachi, Kyoto, Japan). Freeze-dried nanoparticle powders were affixed onto specimen stubs using conductive adhesive. Excess sample was gently removed using an air bulb, and the samples were then gold-coated under vacuum. The observations were performed with an acceleration voltage of 5 kV [35].

2.7.4. X-Ray Diffraction (XRD)

An X-ray diffractometer (D8, Bruker, Karlsruhe, Germany) was used to analyze the freeze-dried NPs’ XRD patterns. The measurement settings were: 45 kV accelerating voltage, 40 mA current, 0.05°/s scanning speed, 5–70° (2θ) range, and 0.02° step size [36].

2.8. Redispersibility Evaluation

The redispersibility was evaluated according to Hu et al. [37]. Nanoparticle suspension (20 mL) was lyophilized and then redispersed in deionized water (20 mL), adjusted to pH 5.5, and stirred (500× g, 2 h). Particle size, PDI, zeta potential, and EE were analyzed according to methods outlined in Section 2.5 and Section 2.6.

2.9. Stability Evaluation

2.9.1. Thermal Stability

The thermal stability was assessed in accordance with Chen et al. [38]. Freshly prepared nanoparticle suspensions (15 mL) were incubated for 2 h at 30, 40, 50, 60, 70, and 80 °C. After cooling, particle size, PDI, and zeta potential were examined according to Section 2.5. Additionally, the AKP retention rate was determined, and the calculation equation was as follows:

$$[eqn]$$

2.9.2. Ionic Stability

The ionic stability was assessed following Chen et al. [39]. Samples (2 mL) were thoroughly mixed with NaCl (2 mL) solutions to final concentrations of 0, 25, 50, 75, 100, and 150 mmol/L. After storing (4 °C, 2 h), particle size, PDI, and zeta potential were assessed as reported in Section 2.5.

2.9.3. pH Stability

The pH stability was evaluated in accordance with Yang et al. [40]. The pH of nanoparticle suspensions was set to 2, 3, 4, 5, 6, 7, and 8. After storing (4 °C, 2 h), particle size, PDI, and zeta potential were evaluated as reported in Section 2.5.

2.9.4. Storage Stability

The storage stability was evaluated following Chen et al. [41]. Freshly prepared suspensions were stored (4 °C, 30 days). Particle size, PDI, and zeta potential were tested every 5 days as reported in Section 2.4. Meanwhile, the AKP retention rate was evaluated as detailed in Section 2.9.1.

2.10. In Vitro Simulated Gastrointestinal Digestion

The freshly prepared sample solution (20 mL, 1.0 mg/mL) was combined with simulated gastric fluid (SGF, 2.0 mg/mL NaCl, 3.2 mg/mL pepsin, pH 2.0) and incubated (37 °C, 200× g, 2 h). Samples were withdrawn every 0.5 h and promptly added to equal volumes of SGF. After stomach digestion, the mixture was neutralized at pH 6.8. Simulated intestinal fluid (SIF, 10 mg/mL bile salts, 1.5 mg/mL trypsin, 6.8 mg/mL KH_2_PO_4_, 8.8 mg/mL NaCl, pH 6.8) was infused into each combination (40 mL) and incubated (37 °C, 200× g, 4 h), sampling every 0.5 h and replenishing with equal volumes of SIF [42]. Particle size and AKP content at each time interval were measured according to Section 2.5 and Section 2.6. The cumulative release rate of AKP was established as follows:

$$[eqn]$$

Release kinetics analysis was performed using the following mathematical models:

$$[eqn]$$ $$[eqn]$$ $$[eqn]$$ $$[eqn]$$

Here, Qt represents the release rate of AKP at time t. K0, K, KH, and Kkp represent the release kinetic rates. n represents the release exponent, and a represents a constant.

2.11. Hypoglycemic Activity

The α-glucosidase, α-amylase, and dipeptidyl peptidase-IV (DPP-IV) were considered core enzymatic targets in studies of the hypoglycaemic activity of food-derived peptides [43]. These enzymes played complementary roles in glycaemic regulation. Therefore, the inhibitory effects of the samples against these three enzymes were simultaneously evaluated to assess their hypoglycemic potential.

2.11.1. α-Glucosidase Inhibition Activity

A sample (50 μL) was blended with α-glucosidase solution (50 μL, 1.26 U/mL) and incubated (37 °C, 10 min). Then, p-nitrophenyl-α-D-glucopyranoside solution (100 μL, 6 mmol/L) was added and incubated (37 °C, 1 h). The reaction was stopped by the addition of Na_2_CO_3_ solution (1 mL, 1 mol/L), and absorbance was measured at 405 nm [42]. The α-glucosidase inhibition rate is calculated using the following equation:

$$[eqn]$$

where A1, A2, A3, and A4 denoted the absorbance values of the sample group (buffer, sample, and enzyme), sample blank group (buffer and sample), blank control group (buffer and enzyme), and blank group (buffer only), respectively.

2.11.2. α-Amylase Inhibition Activity

A sample (50 μL) was combined with α-amylase solution (50 μL, 1.25 U/mL) and incubated (37 °C, 10 min). Then, starch (100 μL, 1%, w/v) was introduced and incubated (37 °C, 10 min). Finally, DNS reagent (400 μL) was incorporated and heated (100 °C, 15 min). Upon cooling, deionized water (1 mL) was added, and the absorbance at 540 nm was analyzed [42]. The α-amylase inhibition rate is calculated by referring to Equation (9) in Section 2.11.1.

2.11.3. DPP-IV Inhibition Activity

In the assay plate, DPP-IV enzyme (20 μL) and the sample (30 μL) were added and mixed, then incubated (37 °C, 10 min). Subsequently, 170 μL of substrate (substrate: buffer = 1:9) was introduced and incubated (37 °C, 30 min). Fluorescence intensity was gauged using a microplate fluorometer (excitation wavelength 360 nm, emission wavelength 460 nm) [44]. Equation to compute DPP-IV inhibition rate:

$$[eqn]$$

where F1, F2, and F3 denote the fluorescence intensity of the blank group (buffer), control group (buffer and enzyme), and sample group (buffer, sample, and enzyme), respectively.

2.12. Data Analysis

All experiments were conducted using three independently prepared samples (n = 3), and each independent sample was measured in triplicate. Statistical analysis was conducted using SPSS 20.0 software, and data were expressed as mean ± standard deviation. Normality of data distribution was assessed using the Shapiro–Wilk test, and homogeneity of variances was evaluated using Levene’s test. Differences were assessed by an independent-samples t-test and one-way ANOVA followed by Duncan’s test. All statistical tests were performed based on the degrees of freedom (df) automatically calculated by the software, with p < 0.05 indicating a statistical difference. Graphs were generated using Origin 2021.

3. Results and Discussion

3.1. Physicochemical Characterization of the AKP

As shown in Table 1, AKP contained a total of 17 amino acids, with moderately hydrophobic amino acids (such as Gly, Ala, Val, Met, Ile, Leu, Phe, and Pro) accounting for approximately 41.75%. This high proportion of hydrophobic amino acids enabled AKP to interact with the hydrophobic regions of OVA through hydrophobic interactions, thereby achieving more embedding within NPs. Additionally, the amino acid composition of AKP was closely related to its potential hypoglycemic activity. Studies demonstrated that high hydrophobicity was an important structural characteristic of potential α-glucosidase and α-amylase inhibitory peptides [45,46]. Such peptides could form strong binding affinities with enzyme active sites, leading to significant inhibitory effects. Most DPP-IV inhibitory peptides were also enriched in hydrophobic amino acids, with Pro, Leu, and Ala being the most common. This may be associated with the binding preference of DPP-IV for hydrophobic amino acids [47]. Yang et al. [48] reported that peptides containing Pro at the N-terminal region usually exhibited strong DPP-IV inhibitory activity. As depicted in Figure 1, the Mw of AKP was mainly distributed in the range of 500–1000 Da. Research reported that peptides with an Mw below 1000 Da exhibited superior hypoglycemic activity [49], suggesting that AKP possessed favorable hypoglycemic potential.

3.2. Particle Size, PDI, Zeta Potential, EE, and LC

The influence of wall material composition on AKP-loaded NPs was presented (Figure 2). Compared to OVA-AKP, CS/OVA-AKP demonstrated a smaller particle size and PDI (mass ratio of CS/OVA was 1:3, 1:2, and 1:1, respectively) (Figure 2a). This may result from the electrostatic attraction between negatively charged OVA and positively charged CS, which increased the intermolecular crosslinking density [18]. However, as the CS content increased, both the particle size and PDI of CS/OVA-AKPs showed an increasing trend. The best stability and dispersibility of the NPs were observed at a 1:2 CS/OVA ratio, where the particle size (292 ± 4 nm) and PDI (0.233 ± 0.036) reached their minimum values. This suggested that, within an appropriate range, the addition of CS strengthened the electrostatic repulsion between NPs, making the nanoparticle structure more compact and favoring the reduction in particle size [50]. Meng et al. [51] found that as the content of carboxymethyl dextrin increased, the particle size of curcumin-loaded zein-carboxymethyl dextrin NPs initially dropped and then rose. The smallest particle size was achieved when the ratio of carboxymethyl dextrin to corn zein was 2:1. To investigate the effect of the wall-material assembly sequences on NPs, OVA/CS-AKP 2:1 was prepared at a 2:1 OVA/CS ratio. The particle size and PDI of OVA/CS-AKP at 2:1 were higher than those of 1:2 (p < 0.05). CS/OVA-AKP formed a composite structure, attributed to electrostatic interactions between the amino groups of CS and the carboxyl groups on the negatively charged OVA [52]. Meanwhile, the CS molecular chains enhanced the crosslinking density through hydrogen bonds and van der Waals forces. This formed a dense interface for the NPs, which prevented aggregation caused by hydrophobic interactions, thus reducing the particle size and improving dispersibility [53]. Previous research has reported that polysaccharides can enhance steric hindrance and electrostatic repulsion between protein-based nanoparticles, thereby improving their colloidal stability [54]. In contrast, for OVA/CS-AKP 2:1, the hydrophobic regions of OVA (such as tryptophan residues) were directly exposed to the aqueous phase, causing partial aggregation of NPs through hydrophobic interactions [17]. Through molecular dynamics simulations, Lee et al. [55] further demonstrated that the exposure of hydrophobic protein residues increased the surface free energy of the NPs, significantly promoting aggregation. The above results indicated that, compared to OVA/CS-AKP 2:1, CS/OVA-AKP 1:2 exhibited better stability and dispersibility.

The repulsive or attractive forces between NPs are important factors affecting the stability of nanoparticle suspensions and are commonly assessed using the zeta potential [56]. As depicted in Figure 2b, OVA-AKP’s zeta potential was −24.8 ± 0.9 mV, mainly due to the carboxyl groups (-COO-) carried by OVA under pH 5.5 conditions (above its isoelectric point, pI = 4.5) [12]. For the CS/OVA-AKPs, the incorporation of CS as the outermost layer changed the zeta potential from negative to positive. This change occurred because the positively charged CS chains tightly adsorbed onto the preformed, negatively charged OVA-AKP surface via electrostatic attraction, forming an integral outer shell that dominated the surface charge properties and resulted in charge reversal from negative to positive. With increasing CS content, the zeta potential increased and then plateaued, indicating a possible saturation in electrostatic interaction between CS and OVA. Zhang et al. [57] found that quercetin-loaded NPs’ zeta potential steadily increased with quaternized chitosan (positively charged). In contrast, CS-AKP showed a significant positive charge (+60.0 ± 2.0 mV), attributable to the protonation of amino groups (-NH_3_^+^) along the CS molecular chain under acidic conditions, conferring a strong positive charge [58]. After adding OVA, the zeta potential of OVA/CS-AKP 2:1 decreased to +32.1 ± 1.0 mV, which was due to the negative charge of OVA neutralizing part of the positive charge on the CS-AKP surface. Furthermore, partial neutralization of surface charge weakened interparticle electrostatic repulsion, while exposed hydrophobic protein sites promoted local aggregation. Consequently, OVA/CS-AKP exhibited a larger particle size and poorer dispersibility than CS/OVA-AKP. This additional discussion further elucidated the intrinsic mechanism by which the wall-material assembly sequences regulated nanoparticle structure and stability through modulation of surface charge composition.

EE and LC are commonly employed to analyze the encapsulation ability of NPs, as presented in Figure 2c. CS/OVA-AKPs exhibited higher EE of AKP compared to OVA-AKP (p < 0.05). This indicated that the single-wall material exhibited a weak binding affinity for AKP, resulting in low loading efficiency or leakage. In contrast, composite-walled could more effectively encapsulate AKP within the NPs through electrostatic interactions [59]. The EE of CS/OVA-AKPs decreased with increasing CS content. This trend aligned with Liu’s findings [60], in which a similar decrease pattern in EE with rising carboxymethyl cellulose content in apigenin-loaded NPs can be observed. Compared to CS/OVA-AKP 1:2 (81.6 ± 1.3%), OVA/CS-AKP 2:1 had a lower EE (75.4 ± 1.3%) (p < 0.05). This difference was likely attributed to stronger interactions among the components in CS/OVA-AKP 1:2, which resulted in the formation of a more compact interfacial layer and network structure. Such a nanoparticle architecture effectively reduced the loss of bioactive compounds during preparation, thereby leading to a higher EE. In summary, the assembly sequences of CS and OVA significantly affected the properties of the NPs. Consequently, CS/OVA-AKP 1:2 and OVA/CS-AKP 2:1 were selected for further study and abbreviated as CS/OVA-AKP and OVA/CS-AKP.

3.3. Structural Analysis

3.3.1. FTIR Analysis

FTIR spectroscopy is commonly applied to recognize the structure of biopolymers and provides information on characteristic absorption peaks and intermolecular interactions [61]. As illustrated in Figure 3, the FTIR spectrum of OVA exhibited characteristic peaks at 3283 cm^−1^, 1640 cm^−1^, 1536 cm^−1^, and 1391 cm^−1^, corresponding to O-H stretching, C=O stretching (amide I band), N-H bending (amide II band), C-N stretching, and N-H bending (amide III band), respectively [62]. The CS spectrum showed an O-H stretching peak at 3288 cm^−1^ and a C-O-C valence vibration peak near 1025 cm^−1^ [63]. AKP mainly included an O-H characteristic peak at 3252 cm^−1^, as well as amide II and III bands at 1560 cm^−1^ and 1390 cm^−1^, respectively. CS/OVA-AKP and OVA/CS-AKP showed O-H peak at 3278 cm^−1^ and 3275 cm^−1^, respectively, which were shifted from individual OVA and CS, suggesting hydrogen bonding between CS amides and OVA hydroxyl groups [64]. This phenomenon was also observed in curcumin-loaded NPs by Zhan et al. [65]. The -CH_2_ stretching peak of OVA at 2924 cm^−1^ weakened post-encapsulation, with blue shifts to 2929 cm^−1^ in both CS/OVA-AKP and OVA/CS-AKP, suggesting hydrophobic interactions between OVA, CS, and AKP [66]. Moreover, after the formation of AKP-loaded NPs (CS/OVA-AKP and OVA/CS-AKP), shifts were observed in the amide I (1640 cm^−1^) and amide II (1536 cm^−1^) bands of OVA, suggesting that electrostatic interactions were present within the system [67]. These results suggested that CS and OVA formed AKP-loaded NPs through electrostatic interactions. This was consistent with Liu et al. [68], where gallic acid-loaded ovalbumin–chitosan NPs showed similar interactions. The formation of CS/OVA-AKP and OVA/CS-AKP was primarily governed by multiple molecular forces, including electrostatic attractions, hydrophobic effects, and hydrogen bonding among OVA, CS, and AKP components.

3.3.2. 3D Fluorescence Spectroscopy Analysis

Endogenous fluorophores (tryptophan, tyrosine, and phenylalanine) in proteins are sensitive to microenvironmental changes [69]. The structural changes of OVA can be displayed through 3D fluorescence spectra [70]. As shown in Figure 4, the strongest fluorescence peak 1 (ex: 282 nm, em: 334 nm) of OVA was mainly attributed to the π → π* transition of tryptophan residues [34]. A secondary peak 2 (ex: 238 nm, em: 332 nm) was likely associated with weak fluorescence from the polypeptide backbone, arising from amide chromophores in peptide bonds via n → π* transitions [71], and/or additional signals arising from heteroaromatic rings [72]. After AKP-loaded NPs formed, the fluorescence intensity of peaks 1 and 2 decreased significantly. During nanoparticle formation, tryptophan or tyrosine residues may have acted as binding sites during encapsulation. This changed the conformation and microenvironment of OVA, causing a quenching effect [73]. Compared to OVA-AKP, the fluorescence intensity of CS/OVA-AKP and OVA/CS-AKP further decreased, suggesting that the complex formation between the composite-walled materials may have also led to fluorescence quenching of OVA. Among them, CS/OVA-AKP showed a more obvious decrease in peak intensity. Compared to OVA, the emission wavelength of peak 1 of AKP-loaded NPs shifted, which could have been caused by conformational changes of OVA during nanoparticle formation. Milião et al. [18] found that electrostatic interactions between chitosan and ovalbumin induced conformational changes in the protein, accompanied by a shift in fluorescence peaks.

3.3.3. SEM Analysis

The microstructure and morphology of AKP-loaded NPs after freeze-drying were observed using SEM. As depicted in Figure 5, OVA-AKP featured a smooth spherical microstructure with aggregation between NPs, possibly due to aggregation occurring between OVA-AKP during the freeze-drying process [64]. CS-AKP displayed a rough surface and irregular shape from CS adsorbing onto AKP, leading to crosslinking interactions between NPs [74]. Compared with OVA-AKP and CS-AKP, NPs encapsulated by the composite-walled materials had a more compact arrangement. CS/OVA-AKP appeared spherical or oval with uniform size and shape. On the other hand, OVA/CS-AKP were tightly packed and showed agglomeration. This could be ascribed to the exposure of hydrophobic amino acids on the outer surface of OVA, triggering hydrophobic aggregation between NPs [75]. These phenomena indicated that the wall-material assembly sequences affected the interactions between NPs and, consequently, the surface characteristics of the NPs.

3.3.4. XRD Analysis

The crystallinity of the main material in NPs and the interactions between biopolymers are frequently assessed using XRD [36]. As shown in Figure 6, AKP exhibited sharp diffraction peaks at 23.1°, 25.7°, 31.9°, and 32.8°, indicating its highly crystalline structure [22]. CS showed a relatively broad diffraction peak at around 19.9°, while OVA displayed two broader peaks at approximately 8.8° and 19.4°, indicating that both individual natural macromolecular carriers had an amorphous structure [76]. In the XRD patterns of AKP-loaded NPs, most of the diffraction peaks of AKP were covered in CS-AKP and OVA-AKP. Weng et al. [77] observed that after curcumin (Cur) was encapsulated in Ophiopogon japonicus protein NPs, its sharp diffraction peaks disappeared, and Cur shifted from a crystalline to an amorphous state. Both CS/OVA-AKP and OVA/CS-AKP showed analogous broad diffraction peaks, with the disappearance of AKP’s diffraction peaks. This indicated that AKP had been successfully encapsulated into the NPs [78]. Amorphous bioactive peptides generally exhibited superior stability and better gastrointestinal absorption compared to their crystalline counterparts [79]. This suggested that AKP encapsulation within CS/OVA matrices may improve its bioavailability.

3.4. Redispersibility

Dried NPs are generally considered advantageous for storage and transportation, offering greater commercial benefits [80]. The freeze-drying process caused partial nanoparticle aggregation, as demonstrated by larger particle sizes, higher PDI, and decreased zeta potential absolute values in re-dispersed NPs compared to freshly prepared NPs (p < 0.05) (Table 2). Moreover, EE decreased in re-dispersed NPs, suggesting that freeze-drying may partially impair encapsulation capability. Liu et al. [60] similarly reported a reduction in zeta potential after redispersion of freeze-dried apigenin-loaded NPs. Compared with peptide-loaded NPs prepared using single-walled materials (OVA-AKP and CS-AKP), those prepared with composite-walled materials (OVA/CS-AKP and CS/OVA-AKP) showed smaller variations in particle size, PDI, zeta potential, and EE following freeze-drying and redispersion. This indicated that the composite wall materials offered enhanced protection for AKP during the lyophilization process, likely due to electrostatic interactions between CS and OVA. Compared to OVA/CS-AKP, CS/OVA-AKP demonstrated smaller changes, likely due to its uniform particle distribution and higher surface charge, effectively preventing aggregation during redispersion [81]. In summary, CS/OVA-AKP exhibited superior redispersibility with minimal particle size increase, the least AKP loss, the highest EE (76.6 ± 1.4%), and optimal dispersion (PDI = 0.272 ± 0.009).

3.5. Stability

3.5.1. Temperature Stability

NPs might undergo high-temperature treatments during manufacturing. As illustrated in Figure 7a, particle size and PDI of CS/OVA-AKP and OVA/CS-AKP slowly increased between 30–60 °C, possibly due to enhanced Brownian motion from heat treatment [82]. At 70–80 °C, particle size and PDI significantly increased (p < 0.05), especially in OVA/CS-AKP, attributed to heat-induced unfolding of OVA structures, exposing hydrophobic groups and promoting aggregation via hydrophobic interactions [83,84]. Compared to OVA/CS-AKP, CS/OVA-AKP exhibited smaller size changes at high temperatures. The absolute value of the zeta potential always remained above 30.0 mV with smaller changes (Figure 7b), indicating better electrostatic repulsion and stability [85]. The content of AKP in NPs (Figure 7c) all decreased with increasing temperature, indicating thermal degradation of AKP [86]. At 80 °C, CS/OVA-AKP exhibited a higher retention rate compared to OVA/CS-AKP (p < 0.05). This may be attributed to the compact structure of CS/OVA-AKP, which partially restricted the denaturation of OVA, thereby providing more effective protection to AKP. In summary, the properties of AKP-loaded NPs were significantly influenced by the temperature, and CS/OVA-AKP showed superior temperature stability.

3.5.2. Ionic Stability

AKP-loaded NPs will encounter various ionic concentration environments during commercial production or within the human digestive tract [87]. Consequently, the particle size, PDI, and zeta potential of AKP-loaded NPs were measured at different salt ion concentrations (0–150 mmol/L), as revealed in Figure 8. The particle sizes of both CS/OVA-AKP and OVA/CS-AKP exhibited an increasing trend with the rise in salt ion concentration. This is due to the aggregation of NPs induced by the introduction of salt ions [88]. CS/OVA-AKP exhibited consistently smaller particle sizes than OVA/CS-AKP (p < 0.05). The results were due to CS adhering to the nanoparticle surface via electrostatic attraction, which enhanced steric hindrance, thereby reducing aggregation caused by the electrostatic shielding effect [89]. Both types of AKP-loaded NPs exhibited good dispersibility in high-salt ion environments (PDI < 0.3), while their zeta potentials decreased with increasing ionic concentration. This was due to the increased ionic strength of the solution, which compressed the double layer on the nanoparticle surface and thereby reduced the electrostatic repulsion between NPs [90]. The zeta potential of CS/OVA-AKP consistently exceeded that of OVA/CS-AKP, and at a salt ion concentration of 75 mmol/L, its zeta potential remained over 30.0 mV, signifying that CS/OVA-AKP exhibited better electrostatic repulsion and superior resistance to the ion shielding effect [85]. In summary, CS/OVA-AKP demonstrated superior ionic stability, and its particle structure can better maintain the stability and dispersibility of the NPs.

3.5.3. pH Stability

AKP-loaded NPs may be exposed to various pH environments during manufacturing, storage, or after ingestion, potentially altering their properties [91]. Therefore, the pH stability of NPs at different pH levels (2–8) was monitored by evaluating their particle size, PDI, and zeta potential, with the results shown in Figure 9. Both NPs showed minor variations in particle size and PDI at pH 2–6. However, these parameters significantly increased at pH 7–8 (p < 0.05), with OVA/CS-AKP exhibiting a more pronounced increase. As the pH rose, the zeta potential of the NPs went down. When the pH approached 6.5, a value at which the protonation degree of CS was significantly reduced (p < 0.05), CS/OVA-AKP tended to aggregate. In contrast, OVA/CS-AKP, due to its surface charge distribution, was more susceptible to pH changes, resulting in more pronounced aggregation [92]. As illustrated in Figure 8b, zeta potentials of both NPs remained relatively stable at pH 2–6 but declined sharply at pH 7–8, particularly for OVA/CS-AKP. This was attributed to the deprotonation of CS, consequently reducing surface charge [93]. Du et al. [94] reported a similar phenomenon, where protein-extended peptide-loaded chitosan-sodium tripolyphosphate NPs exhibited stable particle sizes and zeta potentials at pH 2–5 but showed rapid increases in particle size and significant decreases in zeta potentials at pH 5–6 (p < 0.05). Overall, CS/OVA-AKP demonstrated superior pH stability.

3.5.4. Storage Stability

NPs must maintain durability throughout their shelf life, from commercial production to practical applications [37]. As depicted in Figure 10a, particle size increased gradually with storage time for both types of NPs, likely due to the decreasing absolute zeta potential, which reduced electrostatic repulsion between NPs [95]. Zhu et al. [96] observed a similar trend, reporting a growth of particle size and a reduction in the absolute zeta potential of selenium peptide-loaded lysozyme-xanthan gum NPs over time. Compared to OVA/CS-AKP, CS/OVA-AKP showed less variation in particle size and zeta potential during storage, indicating stronger electrostatic repulsion and a more compact nanoparticle structure, contributing to enhanced stability [97]. Relative to OVA/CS-AKP, CS/OVA-AKP exhibited superior colloidal stability during storage, as demonstrated by minimal fluctuations in hydrodynamic diameter and surface charge. This behavior suggested stronger interparticle repulsive forces and a denser nanostructure, ultimately improving storage durability. Additionally, the AKP retention rate decreased over storage time in both NPs, as demonstrated in Figure 10c. After 30 days, AKP retention rates were 81.48 ± 1.26% for CS/OVA-AKP and 77.71 ± 1.26% for OVA/CS-AKP (p < 0.05), demonstrating superior storage stability of CS/OVA-AKP.

3.6. In Vitro Simulated Gastrointestinal Digestion

The release profile of AKP was shown in Figure 11a. In the simulated gastric environment, free AKP demonstrated a fast release rate, with the cumulative release reaching 76.9 ± 2.7% after 2 h. This result may be due to degradation by the highly acidic gastric juice and the complex enzymatic environment [98]. In contrast, the cumulative release rates of 20.9 ± 2.1% for CS/OVA-AKP and 35.8 ± 2.3% for OVA/CS-AKP were observed during simulated gastric digestion. The higher release rate of AKP from OVA/CS-AKP may be due to the higher surface exposure of OVA in this formulation, which is more susceptible to pepsin digestion than chitosan. Previous studies have reported similar conclusions, suggesting that conformational differences, resulting in varying molecular weight chain flexibility, can affect pepsin accessibility, and that the surface properties of particles can influence digestion properties [99,100]. Following 4 h of incubation in the intestinal environment, free AKP showed a cumulative release rate of 22.2 ± 3.2%, whereas CS/OVA-AKP and OVA/CS-AKP released 61.1 ± 2.1% and 53.0 ± 2.0%, respectively. CS/OVA-AKP exhibited higher intestinal release due to surface-adsorbed CS providing partial resistance to pancreatic enzymatic hydrolysis, allowing more AKP to reach the intestinal environment [101]. Song et al. [102] reported similar findings, indicating enhanced intestinal release of crocin encapsulated in zein NPs coated with chitosan. In addition, Ziebarth et al. [103] reported that when protein-polysaccharide nanoparticles loaded with liraglutide (LIRA) achieved an in vitro cumulative release of approximately 61%, the nanosystem exhibited a hypoglycemic effect comparable to that of subcutaneous LIRA administration in a type II diabetic rat model. This finding suggested that the release extent observed in the present study might possess a certain degree of biological.

Nanoparticle stability and their release behavior within the gastrointestinal tract are affected by particle size [96]. As indicated in Figure 11b, free AKP experienced significant particle size enlargement in gastric conditions due to pepsin-induced aggregation, indicating instability (p < 0.05). In contrast, both AKP-loaded NPs maintained stable particle sizes in gastric fluid, effectively protecting AKP. Particle sizes increased significantly during intestinal digestion (p < 0.05), likely due to pH, ionic strength, and enzymatic conditions. CS/OVA-AKP exhibited a smaller particle size and slower release rates than OVA/CS-AKP during digestion. Guo et al. [104] prepared curcumin and resveratrol-co-loaded NPs and found that smaller NPs exhibited slower release rates during gastrointestinal digestion. Overall, AKP-loaded NPs effectively prevented AKP gastric release and protected it from being damaged by strong acid and pepsin, with CS/OVA-AKP demonstrating superior intestinal targeting.

The release profiles of Free AKP, OVA/CS-AKP, and CS/OVA-AKP were analyzed using four classical drug release kinetic models: zero-order, first-order, Higuchi, and Korsmeyer–Peppas [105]. As shown in Figure 12 and Table 3, the release of free AKP was best described by the first-order model (R^2^ = 0.9866), indicating a concentration-dependent release. In contrast, the Korsmeyer–Peppas model showed the best fit for both OVA/CS-AKP and CS/OVA-AKP. In the Korsmeyer–Peppas model, the magnitude of the release exponent (n) was related to the underlying release mechanism. When n < 0.45, the release followed Fickian diffusion. When 0.45 < n < 0.89, the release corresponded to non-Fickian transport. When n ≥ 0.89, the release behavior was characteristic of Case II transport. The release exponent n for OVA/CS-AKP was 0.7500 (0.45 < n < 0.89), which was consistent with a non-Fickian diffusion mechanism driven by the synergistic effect of drug diffusion and carrier erosion [106]. For CS/OVA-AKP, n was 0.9899 (approaching 1), indicating a Case II transport predominantly controlled by carrier erosion [107]. This could be attributed to the dense outer CS layer, which slowed medium penetration, thereby reducing the burst release of AKP in the in vitro simulated gastric stage and enabling sustained release in the simulated intestinal environment as the CS layer gradually degraded. In summary, compared with free AKP, OVA/CS-AKP and CS/OVA-AKP demonstrated a shift from concentration-dependent release to carrier-controlled release. The assembly sequences of wall materials can modulate the release mechanism of the AKP-loaded NPs, with CS/OVA-AKP exhibiting more desirable sustained-release properties. It should be acknowledged that, despite these observations, this static model provided comparative release profiles rather than predictive bioavailability, and future in vivo validation of the NPs remains necessary.

3.7. Hypoglycemic Activity

The protective effect of NPs on AKP activity was evaluated by determining the hypoglycemic activity of free AKP, AKP-loaded NPs, and OVA-CS NPs (without AKP) before and after simulated gastrointestinal digestion. As shown in Figure 13, free AKP showed inhibition rates of 29.2 ± 1.1% for α-glucosidase, 26.7 ± 1.2% for α-amylase, and 22.0 ± 0.6% for DPP-IV before simulated digestion. In comparison, CS/OVA-AKP (OVA/CS-AKP) exhibited higher inhibition rates of 41.1 ± 1.0% (38.2 ± 1.2%) for α-glucosidase, 37.5 ± 1.5% (35.1 ± 1.5%) for α-amylase, and 36.1 ± 1.2% (32.0 ± 1.0%) for DPP-IV. These results indicated that encapsulation within NPs effectively enhanced AKP’s in vitro hypoglycemic activity. Pan et al. [108] reported similar observations, where curcumin-loaded casein NPs exhibited higher bioactivity compared to free curcumin, attributed to improved dispersion. Notably, CS/OVA-AKP demonstrated better activity performance compared to OVA/CS-AKP. In addition, OVA-CS showed only minimal inhibitory activities against α-glucosidase, α-amylase, and DPP-IV, which were significantly lower than those of AKP-loaded NPs (p < 0.05), indicating a negligible contribution from the carrier matrix.

After simulated digestion, inhibition rates for α-glucosidase (13.9 ± 1.3%), α-amylase (9.7 ± 1.4%), and DPP-IV (8.8 ± 0.8%) significantly decreased (p < 0.05) for free AKP due to degradation by acidic gastric conditions and digestive enzymes [109]. For α-glucosidase, α-amylase, and DPP-IV, the inhibition rates by CS/OVA-AKP were 41.1 ± 1.0%, 27.0 ± 1.5%, and 24.0 ± 1.5%, respectively, compared to 38.2 ± 1.2%, 21.6 ± 1.7%, and 19.6 ± 1.5% for OVA/CS-AKP. Although the hypoglycemic activity decreased post-digestion, AKP-loaded NPs remained higher (p < 0.05) than that of free AKP, likely due to the nanoparticle delivery system, which can effectively inhibit the degradation of AKP during digestion [110]. Among them, CS/OVA-AKP exhibited higher α-glucosidase, α-amylase, and DPP-IV inhibitory effects before and after simulated digestion (p < 0.05). However, these results merely reflected the potential in vitro hypoglycemic activity of these peptide-loaded NPs, and the in vivo absorption and therapeutic equivalence remain to be further clarified. Future studies should use cell models and animal experiments to verify their efficacy and mechanism of action in biological systems.

3.8. The Formation Process of AKP-Loaded NPs

The formation mechanisms of CS/OVA-AKP and OVA/CS-AKP are shown in Figure 14. In CS/OVA-AKP, OVA initially formed a negatively charged complex with AKP through hydrogen bonds and hydrophobic interactions. Positively charged CS then coated the complex, self-assembling into a complete CS/OVA composite wall material. This structure trapped the unstable protein chains inside the NPs, while the outer CS layer extended into a brush-like or network structure, preventing nanoparticle aggregation and enhancing NPs’ stability in both the environment and the gastrointestinal tract [54]. FTIR characteristic peak shifts, more pronounced OVA fluorescence quenching, and uniform spherical morphology in SEM collectively confirmed this mechanism. In contrast, in OVA/CS-AKP, CS and AKP were combined via electrostatic interactions, reducing the surface charge. OVA weakly adsorbed to the surface, making it difficult to form a dense wall material. The rigid CS layer caused a loose internal structure, and the OVA exposed hydrophobic sites, which promoted aggregation between NPs [17]. This was reflected in weaker fluorescence quenching and significant aggregation in SEM, resulting in poorer stability. Although XRD showed AKP in an amorphous form in both systems, the differences in wall-material assembly explained the superior encapsulation efficiency, stability, and delivery performance of CS/OVA-AKP. Additionally, the outer polysaccharide layer may have reduced OVA’s allergenic potential, offering greater application potential.

4. Conclusions

This study confirmed that the wall-material assembly sequences notably influenced the structure and properties of AKP-loaded NPs. Compared to OVA/CS-AKP, CS/OVA-AKP exhibited smaller particle size, better dispersibility, and higher EE. The formation of CS/OVA-AKP primarily relied on electrostatic interactions, hydrogen bonding, and hydrophobic forces, resulting in a dense nanoparticle structure with more uniform morphology. In contrast, OVA/CS-AKP exhibited more pronounced aggregation, likely owing to the increased exposure of hydrophobic regions in OVA. Furthermore, CS/OVA-AKP displayed superior resistance to destabilization factors, including thermal treatment, salt concentration, pH, and storage duration. This better stability can be attributed to the cationic nature of CS, the electrostatic repulsion, and the steric hindrance effects provided by the rigid polysaccharide chain of CS. In simulated gastrointestinal digestion, compared with OVA/CS-AKP, CS/OVA-AKP exhibited a 14.97% reduction in cumulative AKP release in the simulated gastric phase, while AKP release in the simulated intestinal phase increased by approximately 8.14%, indicating improved sustained release. Moreover, the in vitro hypoglycemic activity of encapsulated AKP was improved both before and after simulated digestion, with CS/OVA-AKP showing the best in vitro hypoglycemic activity. Nevertheless, these results indicate a potential hypoglycemic function, and its actual therapeutic equivalence or in vivo efficacy still requires further investigation in future studies. In conclusion, the assembly sequences of the wall materials are crucial for developing a delivery system to enhance the stability of bioactive peptides and protect their bioactivity.