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7 February 2026

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Active Biological Film Improves the Quality of Mutton During Super-Chilling Storage: Effects on Myofibrillar Protein Characteristics and Physicochemical Properties

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College of Food Science and Engineering, Shanxi Agricultural University, Jinzhong 030801, China
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Foods2026, 15(4), 609;https://doi.org/10.3390/foods15040609
This article belongs to the Section Meat
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Abstract

Preventing spoilage in food products, particularly in those highly susceptible to rapid deterioration like mutton, has been a persistent challenge in the food industry. In this study, an Active Biological Film (ABF) was developed using chitosan (CS) and whey protein isolate (WPI), with the addition of 0.01 wt% titanium dioxide (TiO2) and 0.1 wt% white pepper essential oil (WPEO). This ABF was applied to preserve fresh mutton at super-chilling temperatures of −1.7 ± 0.2 °C. The effects of ABF on myofibrillar protein (MP) oxidation and structural characteristics, as well as on the microbial status, physicochemical properties, and sensory quality of mutton, were systematically evaluated. The results demonstrated that, compared to the control group (CK), ABF treatment significantly enhanced the total sulfhydryl content, protein solubility, and zeta potential of MPs, while reducing carbonyl content, surface hydrophobicity, and particle size. MPs in the ABF group showed a higher α-helix proportion and a lower random coil content, along with a notable increase in intrinsic fluorescence intensity. Scanning electron microscopy (SEM) revealed a denser gel structure. Additionally, ABF effectively inhibited microbial growth in mutton, delayed pH increase, reduced thiobarbituric acid-reactive substances (TBARS) and total volatile basic nitrogen (TVB-N), and improved sensory scores, extending mutton shelf life by over 10 days. Therefore, the ABF effectively inhibited oxidation in MPs, maintained their structural integrity, and preserved mutton quality during super-chilling storage.

1. Introduction

Mutton, a valuable source of animal protein, is favored by consumers for its high nutritional value [1]. However, mutton is prone to quality deterioration during storage and distribution due to microbial growth, enzymatic reactions, and oxidative reactions.
Such quality deterioration—manifested as color fading, drip loss, oxidation of lipids and proteins, and off-flavor formation—impairs product quality and constrains the long-term development of mutton production systems [2,3]. Among these factors, oxidative denaturation of muscle proteins, especially myofibrillar proteins (MPs), is a major contributor to meat-quality deterioration [4]. Oxidation of MPs during storage leads to nutritional losses and reduced functional properties [5,6].
To delay spoilage and extend its shelf life, low-temperature storage is one of the most widely used techniques. Super-chilling is a low-temperature preservation technology that stores fresh food at temperatures below 0 °C but above its freezing point, ensuring that the food remains in a non-frozen state. This method helps avoid structural damage and excessive drip loss caused by freezing, thereby extending food freshness [7]. Lin Zhang et al. [8] analyzed the changes in physicochemical properties and microbial indicators of surimi products under super-chilling (−1 °C) and refrigeration (4 °C) conditions, and found that super-chilling significantly inhibited microbial metabolism and lipid/protein oxidation. Ji et al. [9] assessed the physicochemical properties of chicken breast muscle at −1.5 °C (super-chilling) and 4 °C (refrigeration), and reported that −1.5 °C storage mitigated decreases in muscle solubility and in total and reactive sulfhydryl contents.
However, the effectiveness of a single physical preservation method is inherently limited, as it often fails to comprehensively address complex, interrelated spoilage pathways in meat products (e.g., microbial growth, lipid oxidation, and protein degradation) [10,11]. Therefore, combining physical preservation with complementary approaches (e.g., active packaging, natural antimicrobials) to achieve synergistic preservation effects has become a necessary and promising strategy [12]. Active packaging, a smart packaging strategy, incorporates functional substances with antimicrobial and antioxidant properties into materials, allowing controlled interactions with the food and its surrounding environment to preserve and improve quality. This approach has become a key focus in food packaging research [13,14]. Biopolymer-based active packaging films, particularly composite films made from natural biodegradable materials such as whey protein isolate (WPI) and chitosan (CS), have received increasing interest because of their good compatibility with food matrices and their suitability for forming functional packaging structures [15]. By incorporating functional ingredients, these films can be endowed with enhanced bioactivity. Owing to its photocatalytic activity, titanium dioxide (TiO2) has been applied in biopolymer films to confer protective functions against ultraviolet radiation, microbial growth, and oxidative degradation [16,17]. Essential oils, rich in terpenes and other aromatic compounds, exhibit strong antimicrobial and antioxidant activities, thereby enhancing the functionality of active films [18,19]. In previous experiments, we synergistically incorporated TiO2 and white pepper essential oil (WPEO) synergistically into the CS/WPI composite matrix to develop an efficient active packaging material [20]. However, systematic research on its impact on protein stability and overall mutton quality during super-chilling storage remains limited.
Based on this, the present study applied a CS/WPI bio-composite, freshness-preserving active film containing 0.01 wt% TiO2 and 0.1 wt% WPEO to fresh mutton. The samples were stored under super-chilling conditions (−1.7 ± 0.2 °C) to evaluate the preservation performance of the film. This study investigated the effects of the bio-based active film on MP oxidation (total sulfhydryl content, carbonyl content, surface hydrophobicity, and protein solubility) and on the structural properties (zeta potential, particle size, intrinsic fluorescence, and gel microstructure) of mutton myofibrillar proteins (MPs) over 30 days of storage. Additionally, the study systematically evaluates its effects on mutton microbial indicators, physicochemical properties (pH, TBARS, TVB-N), and sensory quality. These results are expected to provide a theoretical basis for applying this active packaging material to chilled mutton preservation.

2. Material and Methods

2.1. Material and Reagents

Super-chilled mutton samples were acquired from Shanxi Biansai Sheep Breeding Technology Service Co., Ltd. (Datong, China). Chitosan (deacetylation degree of 95%) and nano-TiO2 (particle size of 5–10 nm, purity of ≥99.8%) were purchased from Shanghai Acmec Biochemical Co., Ltd. (Shanghai, China). WPI (protein content of >90 wt%) was obtained from Beijing Jialikangyuan International Trade Co., Ltd. (Beijing, China). White pepper essential oil (WPEO) was provided by Tianjin Chunhe Technology Development Co., Ltd. (Tianjin, China). All other reagents were of analytical grade and obtained from Beijing Solab Technology Co., Ltd. (Beijing, China).

2.2. Preparation of Active Biodegradable Film (ABF)

The ABF was prepared according to Wang et al. [20]. Briefly, chitosan (CS, 1% w/v) was dissolved in 1% (v/v) acetic acid under continuous stirring and then heated at 60 °C for 40 min. Whey protein isolate (WPI, 0.4 g) was dissolved in 100 mL of distilled water, adjusted to pH 8.0, and heated at 85 °C for 30 min with constant stirring to obtain heat-denatured WPI. The CS solution was then mixed with the WPI solution, followed by the addition of glycerol (1% w/v) and stirring for 1 h. TiO2 particles were added to the mixture and homogenized using a homogenizer (JH-ZS1, Hangzhou Jinghao Machinery Co., Ltd., Hangzhou, China) at 1000 r/min for 10 min, followed by ultrasonication using an ultrasonic device (JY92-IIDN, Ningbo Xinyi Ultrasonic Equipment Co., Ltd., Ningbo, China) for 20 min. Subsequently, WPEO (0.1% w/v) was added and the solution was homogenized for an additional 10 min. The resulting film-forming solution was cast onto an acrylic plate (12 cm × 12 cm) and dried in an oven at 25 ± 2 °C for 24 h. After drying, the thickness of the ABF was measured to be 0.024 nm.

2.3. Preparation of Mutton Samples and Treatments

Ten castrated male Shanxi Small-Tailed Han sheep (36 months old) with similar body weights were used in this study and were raised under identical feeding conditions. All animals were slaughtered under standard commercial conditions by Shanxi Biansai Sheep Breeding Technology Service Co., Ltd. The carcasses were obtained within 24 h postmortem and transported to the laboratory at 4 °C. The longissimus dorsi muscle (located along the dorsal region adjacent to the spine) was excised and used for subsequent analyses. After removing excess fat, the muscles were cut into 50 ± 5 g pieces under sterile conditions.
Samples were randomly assigned to two groups: an ABF-treated group, in which the ABF was placed directly between the mutton sample and the shrink-wrap film to ensure contact, and an untreated control group (CK) without ABF. All samples were then packaged in a shrink-wrap film (HYCD-205, Qingdao Haier Refrigerator Co., Ltd., Qingdao, China). Due to the freezing point of mutton being measured as −2 °C in our preliminary experiments, the super-chilling storage temperature was set to (−1.7 ± 0.2) °C, and all packaged samples were stored at this temperature for 30 days. Samples were collected at 0, 1, 4, 7, 10, 15, 20, 25, and 30 days for the determination of storage quality indicators. Additionally, sampling was conducted on days 0, 5, 10, 15, 20, 25, and 30 to evaluate changes in MP characteristics.

2.4. Extraction of Myofibrillar Proteins (MPs)

MPs were isolated according to a previously reported protocol [21]. In brief, visible fat and connective tissue were removed from the samples before mincing and weighing. A 5.00 g portion of minced meat was placed into a 100 mL pre-chilled centrifuge tube, and 20 mL of pre-chilled protein extraction buffer (KCl 7.455 g/L, MgCl2 0.4066 g/L, Na2HPO4 3.5814 g/L; pH adjusted to 7.0; ionic strength 0.19 mol/L) was added. The mixture was homogenized for 30 s using a homogenizer (JH-ZS1, Hangzhou Jinghao Machinery Co., Ltd., Hangzhou, China). All procedures were conducted in an ice bath to prevent protein oxidation and denaturation. The homogenate was centrifuged at 2000× g for 15 min at 2 °C, and the supernatant was discarded; the “extraction–homogenization–centrifugation” step was repeated twice (three cycles in total). The resulting pellet was resuspended in 40 mL of pre-chilled washing buffer (NaCl 5.844 g/L; ionic strength 0.10 mol/L), homogenized for 30 s, and centrifuged at 2000× g for 15 min at 2 °C; the supernatant was discarded. Subsequently, the pellet was homogenized again in 40 mL pre-chilled washing buffer for 30 s. After filtration through two layers of gauze, the filtrate was adjusted to pH 6.2 using pre-chilled 0.1 mol/L HCl, followed by centrifugation at 2000× g for 15 min at 2 °C. The supernatant was discarded, and the pellet was collected as myofibrillar proteins. The isolated myofibrillar proteins were stored at 4 °C and analyzed within 48 h.

2.5. Determination of Physicochemical Properties of MPs

2.5.1. Carbonyl Content

The carbonyl content of MPs was quantified using a previously reported DNPH-based method [22]. Briefly, the MPs suspension was diluted to 25 mg/mL, and 0.1 mL of the sample was allowed to react with 2 mL of 2,4-dinitrophenylhydrazine dissolved in 2 mol/L hydrochloric acid, while the control was treated with hydrochloric acid alone. After incubation in an ice bath for 60 min with periodic agitation, protein precipitation was induced using 20% (w/v) trichloroacetic acid. The resulting precipitate was collected by centrifugation (11,000× g, 4 °C, 3 min), rinsed three times with an ethanol/ethyl acetate mixture (1:1, v/v), and air-dried. The pellet was then solubilized in 6 mol/L guanidine hydrochloride and incubated at 50 °C for 45 min. Absorbance was recorded at 280 nm and 370 nm using a Cary 4000 spectrophotometer (Agilent Technologies, Inc., Santa Clara, CA, USA). Carbonyl content was calculated using a molar extinction coefficient of 2.2 × 104 L·mol−1·cm−1 according to the equation below:
Carbonyl   content   nmol / L = A 370 × 10 6 2.2 × 10 4 × C × 10 3
where A370 represents absorbance at 370 nm and C denotes protein concentration (mg/L) determined at 280 nm.

2.5.2. Total Sulfhydryl Content

The total sulfhydryl content of MPs was determined based on the method reported by Kong et al. [23] using 5,5′-dithiobis-(2-nitrobenzoic acid) (DTNB), with minor modifications. An aliquot of MP solution (0.5 mL) was combined with 2.5 mL of Tris–glycine–urea buffer and 20 μL of freshly prepared DTNB solution (0.004 g/mL), with distilled water serving as the blank control. Following a 25 min reaction under light-protected conditions at ambient temperature, absorbance was recorded at 412 nm with a Cary 4000 UV-visible spectrophotometer (Agilent Technologies, Inc., Santa Clara, CA, USA). Total sulfhydryl content was determined according to the equation below:
Total   sulfhydryl nmol / mg = A 412 × 10 6 1.36 × 10 4 × C
where A412 represents the absorbance at 412 nm, 1.36 × 104 L·mol−1·cm−1 is the molar extinction coefficient of DTNB, and C denotes the protein concentration (mg/mL).

2.5.3. Solubility

The solubility of MPs was evaluated using a reported procedure [24]. MPs were diluted to 5 mg/mL with PIPES buffer (0.6 mol/L NaCl, 20 mmol/L PIPES, and 10 mmol/L sodium pyrophosphate; pH 6.2), maintained at 2 °C for 4 h, and then centrifuged at 5000× g for 15 min. The protein concentration of the resulting supernatant was quantified by the biuret assay, and solubility was calculated as the proportion of soluble protein relative to the initial protein content.

2.5.4. Surface Hydrophobicity

The surface hydrophobicity of MPs was assessed using a bromophenol blue (BPB) binding assay based on a reported protocol [25]. MPs were diluted to 5 mg/mL, after which 1 mL of the protein solution was combined with 200 μL of BPB (1 mg/mL). Phosphate buffer was used in place of protein for the blank determination. After incubation at room temperature for 10 min, samples were centrifuged at 2000× g for 15 min. The supernatant was diluted tenfold, and absorbance was recorded at 595 nm using a Cary 4000 spectrophotometer (Agilent Technologies, Inc., Santa Clara, CA, USA). Surface hydrophobicity was expressed as bound BPB, calculated as the difference between total and free BPB.

2.6. Determination of Structural Properties of MPs

2.6.1. Zeta Potential and Particle Size

The hydrodynamic diameter and surface charge of MPs were analyzed at 25 °C using a Zetasizer Nano ZS90 (Malvern Instruments, Malvern, UK) with a 1 mg/mL protein suspension. Particle size was obtained after 1 min of thermal equilibration, whereas zeta potential measurements were conducted following a 2 min equilibration period.

2.6.2. Intrinsic Fluorescence

Intrinsic fluorescence spectra of MPs were recorded at a protein concentration of 0.25 mg/mL using an F-7100 fluorescence spectrophotometer (Hitachi High-Tech Co., Ltd., Tokyo, Japan), following a previously reported protocol [26].

2.6.3. Secondary Structure Analysis

The secondary structure was analyzed following the method described by Luo et al. [27]. Potassium bromide was mixed with the freeze-dried MP sample in a 2:100 ratio, then compressed into a pellet and analyzed using a Tensor 27 Fourier-transform infrared spectrometer (Bruker Co., Ltd., Ettlingen, Germany). Spectra were collected in the range of 4000–400 cm−1 with a spectral resolution of 4 cm−1. Spectra in the amide I region (1600–1700 cm−1) were extracted and analyzed using PeakFit 4.11 software (Systat Software Inc., San Jose, CA, USA).

2.6.4. Microstructure Analysis of MP Gels

A blade was used to cut the MP gels into 3 mm × 3 mm × 2 mm pieces. Samples were fixed overnight at 4 °C in 2.5% glutaraldehyde, followed by three washes with phosphate buffer (pH 7.2). Following this, the samples were gradient-eluted with ethanol of varying concentrations. After the washing process, the samples were vacuum freeze-dried, and microstructural features were examined by scanning electron microscopy (FEI Quanta 450, FEI Company, Hillsboro, OR, USA).

2.7. Determination of Quality Indicators for Mutton

2.7.1. Total Viable Count (TVC)

TVC was quantified by the plate count technique. Minced mutton samples (10 g) were blended with 90 mL of sterile 0.85% saline to prepare the initial dilution, followed by successive decimal dilutions. Appropriate dilutions were spread onto plate count agar (Qingdao Haibo Biotechnology Co., Ltd., Qingdao, China) and incubated at 37 °C for 48 h. Microbial levels were reported as log CFU/g.

2.7.2. pH

The pH was measured following a previously reported procedure [28]. Briefly, 1 g of sample was mixed with 10 mL of distilled water and homogenized for about 30 s using a FA25 high-shear disperser (Shanghai Fluke Fluid Machinery Manufacturing Co., Ltd., Shanghai, China). The pH was determined using an ST2100 pH meter (Ohaus Instruments Co., Ltd., Shanghai, China).

2.7.3. Thiobarbituric Acid-Reactive Substances (TBARS)

Lipid oxidation was evaluated using the TBARS assay following Cheng et al. [29]. Briefly, 5 g of finely chopped mutton was extracted with 15 mL of 7.5% (w/v) trichloroacetic acid supplemented with 0.1% butylated hydroxyanisole and 0.1% ethylenediaminetetraacetic acid. After filtration, 2.5 mL of the extract was mixed with an equal volume of 0.02 mol/L thiobarbituric acid and heated in a boiling-water bath for 40 min. The reaction mixture was subsequently cooled, partitioned with chloroform, and centrifuged at 2000× g for 10 min at 2 °C. The absorbance of the upper phase was recorded at 532 nm using a Cary 4000 spectrophotometer (Agilent Technologies, Inc., Santa Clara, CA, USA).

2.7.4. Total Volatile Basic Nitrogen (TVB-N)

The TVB-N content of mutton was quantified following an established protocol [30]. Briefly, 5 g of chopped meat was dispersed in 50 mL of distilled water, agitated for 30 min, and clarified by filtration. A 5 mL portion of the filtrate (distilled water served as the blank) was combined with 5 mL of a 1% (w/v) magnesium oxide suspension and subjected to steam distillation. Volatile alkaline compounds were collected in 10 mL of 2% (w/v) boric acid and determined by titration with 0.01 mol/L hydrochloric acid.

2.7.5. Sensory Evaluation

The sensory evaluation was conducted using an assessment protocol based on previously published work by Sayadi, Mojaddar Langroodi, et al. [31]. A panel of 10 trained panelists (5 males and 5 females) was selected for the assessment. Prior to formal evaluation, all panelists received a training session for 30 min to familiarize them with the scoring criteria and sensory attributes. The training was considered complete when the panelists were familiar with the evaluation procedure and the difference between each individual score and the panel mean was within one unit. All samples were evaluated by each panelist within a single evaluation session. Odor, color, texture, and overall acceptability were assessed using a hedonic scoring system with nine categories. Lower scores (1–3) corresponded to unsatisfactory sensory quality, mid-range scores (4–6) indicated acceptable attributes, and higher scores (7–9) reflected superior sensory acceptance. Sensory evaluation was conducted in a designated sensory evaluation area under natural light at room temperature (25 ± 2 °C).

2.8. Statistical Analyses

All measurements were performed in triplicate. Data were statistically processed using SPSS software (version 21.0, IBM Corp., North Castle, NY, USA). One-way ANOVA was applied to evaluate differences among groups, and mean separations were performed using Duncan’s multiple range test at a significance level of p < 0.05. Figures were prepared with Origin 2025 (OriginLab Corporation, Northampton, MA, USA).

3. Results and Discussion

3.1. Changes in Oxidation and Structural Properties of MPs

3.1.1. Oxidation Properties of MPs

During storage, protein oxidation driven by hydroxyl-derived reactive species induces modifications of amino acid residues and peptide structures, thereby increasing carbonyl formation; thus, carbonyl content is widely regarded as a reliable indicator of protein oxidation [32]. As shown in Figure 1A, the carbonyl levels of MPs in both groups increased progressively throughout the super-chilling period. At day 0, the carbonyl content of MPs was 1.81 × 10−3 nmol/mg; by day 30, the ABF-treated samples (6.38 × 10−3 nmol/mg) showed significantly lower values than the CK group (8.87 × 10−3 nmol/mg) (p < 0.05), indicating that the ABF effectively slowed protein carbonylation. This inhibitory effect is likely attributed to the white pepper essential oil incorporated in the ABF, as its constituents possess strong radical-scavenging and metal-chelating capacities. These properties inhibit carbonyl generation resulting from the oxidation of amino acid residues, including threonine, proline, arginine, and lysine, thereby reducing MP aggregation and preserving functional characteristics [33]. Consistent findings were reported by Karimi et al. [34], who found that ostrich meat coated with a Malva neglecta mucilage (MLM) nanocomposite containing 0.8% Myrtus communis essential oil (MEO) exhibited markedly lower carbonyl levels at the end of storage (1.74 nmol/mg) than the control group (3.17 nmol/mg).
Figure 1. Effects of ABF on the physicochemical properties of MPs in mutton during super-chilling storage. (A) Carbonyl content; (B) total SH content; (C) solubility; (D) surface hydrophobicity. CK: Control group; CS/WPI/TiO2 + WPEO: Chitosan/Whey protein isolate/TiO2 + White pepper essential oil. Different uppercase letters indicate significant differences (p < 0.05) at the same storage time. Different lowercase letters indicate significant differences (p < 0.05) within the same treatment.
Sulfhydryl groups are widely present in proteins and are prone to oxidation, forming disulfide bonds. Therefore, the total sulfhydryl content is a reliable indicator of protein oxidation [35,36]. As shown in Figure 1B, the total sulfhydryl content of MPs reached a maximum value of 94.37 nmol/mg at day 0 and then declined significantly with prolonged storage (p < 0.05). By day 30, the total sulfhydryl content in the CK group dropped to 71.51 nmol/mg, which is 75.78% of the initial value, while the ABF group retained a higher sulfhydryl content of 78.87 nmol/mg, which is 83.58% of the initial value. This phenomenon is due to the active compounds in white pepper essential oil in the ABF, which adsorb onto sulfhydryl groups and compete for free radicals, inhibiting sulfhydryl oxidation [37]. Similarly, Lan et al. [38] found that a chitosan/polyvinyl alcohol (CS/PVA) active film with ginger essential oil effectively protected sulfhydryl groups throughout seabass storage. Moreover, J. Cheng et al. [39] and Khoder et al. [40] demonstrated that a range of plant-derived compounds, including cinnamaldehyde, rutin, quercetin, and other phytochemicals, preferentially neutralize free radicals, thereby protecting sulfhydryl groups from oxidative damage.
The decline in protein solubility is closely correlated with the extent of protein oxidative denaturation, with higher oxidation levels typically resulting in lower solubility [41]. Figure 1C shows that MP solubility in all samples decreased progressively during super-chilling storage. By day 30, the solubility of MPs in the CK group had dropped from 92.54% to 60.64%. Previous studies have attributed such reductions to conformational and structural alterations induced by oxidation [40]. During storage, disruption of the native MP structure exposes buried sulfhydryl groups, leading to disulfide bond formation, protein cross-linking, aggregation, and reduced solubility [42]. In contrast, the essential oil components and chitosan present in the ABF exerted protective effects on MP structural integrity. Consequently, the ABF treatment maintained a significantly higher MP solubility (68.37%) at day 30 relative to the control samples (p < 0.05). Similar findings were reported by Li et al. [43], who observed that an active coating composed of locust bean gum and sodium alginate enriched with Lippia citriodora Kunth essential oil effectively mitigated the decline in protein solubility in large yellow croaker.
The binding of bromophenol blue (BPB) to proteins reflects the extent of MP unfolding and hydrophobic amino acid residue exposure. Therefore, it serves as an indicator of protein surface hydrophobicity [40]. Figure 1D shows the binding of MPs to BPB over time. At day 0, the MPs exhibited the lowest binding to BPB (5.32 μg). During storage, oxidation increased the binding of proteins to BPB. During the initial storage period (day 0–5), BPB binding showed no statistical difference between the CK and ABF treatments (p > 0.05). However, a pronounced increase in BPB binding was observed in the CK samples at later stages. At day 30, BPB binding in the CK group reached 35.56 μg, whereas a significantly lower value of 27.71 μg was recorded for the ABF group (p < 0.05). This difference is likely associated with the antioxidant activity of the essential oil, which mitigated protein oxidation, limited MP structural unfolding, and reduced the exposure of hydrophobic residues, thereby suppressing the increase in MP surface hydrophobicity [44]. Based on the above analysis, ABF effectively suppressed MP oxidation, likely due to its protective effect on their structure.

3.1.2. Structural Properties of MPs

The zeta potential reflects the surface charge of proteins and is used to assess electrostatic interactions among charged particles [45]. A negative zeta potential is generally observed when negatively charged amino acid residues predominate over positively charged residues in the protein system. As shown in Figure 2A, all samples exhibited a negative zeta potential, indicating that MP surfaces carried net negative charges. This is primarily attributable to deprotonation of acidic residues (e.g., glutamic acid and aspartic acid), which contributes to an overall negative surface charge [46]. Initially, the absolute zeta potential of MPs was 20.33 mV, but it gradually decreased during storage. The decrease was more pronounced in the CK group, reaching 7.08 mV at day 30, while the ABF group maintained a significantly higher absolute zeta potential (9.92 mV) (p < 0.05). Figure 2B–E shows the average particle size of MPs in the mutton samples. On day 0, the average particle size was 164.33 nm. As storage time increased, the particle size significantly increased (p < 0.05), reaching 295.38 nm in the CK group and 253.74 nm in the ABF group at day 30. During storage, disruption of non-covalent interactions promotes protein denaturation and aggregation, which typically reduces the magnitude of the zeta potential and increases MP particle size [41]. In contrast, ABF-treated MPs maintained a more uniform particle-size distribution and a relatively higher zeta potential, indicating improved colloidal stability and suppressed aggregation.
Figure 2. Effects of ABF on the structural properties of MPs in mutton during super-chilling storage. (A–E) Zeta potential and average particle size; (F,G) fluorescence intensity. CK: Control group; CS/WPI/TiO2 + WPEO: Chitosan/Whey protein isolate/TiO2 + White pepper essential oil. Different uppercase letters indicate significant differences (p < 0.05) at the same storage time. Different lowercase letters indicate significant differences (p < 0.05) within the same treatment.
Figure 2F,G present the intrinsic fluorescence spectra of MPs, which can be used to assess the integrity of their tertiary structure. Fresh mutton MPs exhibited the highest fluorescence intensity. Over the storage period, the fluorescence intensity of MPs in both groups gradually decreased. By day 30, the fluorescence intensity in the ABF group decreased more slowly than in the CK group (Figure 2G), indicating that ABF effectively preserved the stability of the MP tertiary structure.
The secondary structure of proteins includes α-helices, β-turns, β-sheets, and random coils, among other elements [47] As shown in Table 1, fresh mutton MPs contained α-helices (38.21%), β-sheets (29.93%), β-turns (20.15%), and random coils (11.71%). Over time, α-helix and β-sheet contents in both the CK and ABF groups decreased, whereas β-turn and random coil contents increased. By the end of storage, α-helix and β-sheet contents in the CK group decreased to 32.09% and 25.07%, respectively, whereas the ABF group maintained higher values of 36.28% and 27.44%, markedly greater than the CK group (p < 0.05). Random coil content in the CK group increased from 11.71% to 17.87%, whereas the ABF group increased to 15.47%. α-helices and β-sheets are typically stabilized by hydrogen bonds formed between carbonyl and amide groups on the peptide chain. When their proportions decrease, hydrogen bonding within the backbone is disrupted, leading to reduced secondary-structure stability [38]. This change shifts the structure from α-helix to random coil, increasing the proportion of random coils [43]. Consistent findings were found by Xie Y et al. [37], who observed that MPs in sea bass treated with a 2% jasmine essential oil active coating showed higher α-helix and β-sheet content than the control group.
Table 1. Effects of ABF on the secondary structure of MPs in mutton during super-chilling storage.
The microstructure of MP gels, including the compactness of its three-dimensional network structure and pore size, is a key indicator for assessing gel functionality [48], observable through scanning electron microscopy (SEM). Figure 3 shows significant differences in the MP gel structure among the samples. MP gels from fresh mutton exhibited a dense structure with small pores. As storage time increased, the microstructure changed significantly. Compared to fresh samples, the CK group exhibited larger voids and a rougher, more porous structure, which became more pronounced as storage time progressed, with the most noticeable changes occurring at day 30. In contrast, the ABF group maintained a more orderly structure with a gel network containing fewer pores, and even at day 30, the gel surface remained relatively continuous and dense.
Figure 3. Effects of ABF on the microstructure of MP gels in mutton during super-chilling storage. CK: Control group; CS/WPI/TiO2 + WPEO: Chitosan/Whey protein isolate/TiO2 + White pepper essential oil.
These structural differences are primarily due to varying degrees of protein oxidation. Oxidation causes denaturation of myosin and actin, inhibiting some crosslinking reactions between amino acid side chains, which leads to a decrease in the gel network’s viscoelasticity, resulting in a looser structure and the formation of aggregates [21,49]. The ABF’s antioxidant activity effectively slowed the oxidation process, preserving the protein structure and maintaining the stability of the gel network [50]. These results further support the positive role of ABF in inhibiting protein oxidation and preserving gel quality. Additionally, Chang et al. [51] found that Gnaphalium affine extract (GAE) at 0.2 mg/g protein exhibited strong free radical scavenging ability, preventing oxidation and protecting protein side chain structures, thereby maintaining the gel network’s integrity. This mechanism is consistent with the effects observed in this study for the composite film.

3.2. Changes in Mutton Quality

3.2.1. TVC

Microbial proliferation is a primary driver of food spoilage, and previous studies have indicated that raw meat is generally regarded as spoiled once the TVC reaches 6.0 log CFU/g [52]. At the beginning of storage, mutton exhibited a TVC of 3.82 log CFU/g (Figure 4A). Typically, the initial TVC of mutton ranges from 102 to 104 CFU, which depends on the animal’s breed, living environment, and slaughtering conditions [53]. Over the course of storage, the TVC of the CK group increased rapidly, reaching 5.83 log CFU/g at day 20, which was below 6 log CFU/g and within an acceptable range. However, by day 25, the TVC reached 6.25 log CFU/g, indicating that the mutton had spoiled. In contrast, the ABF group maintained a TVC of 5.31 log CFU/g at day 30, still within the microbial shelf life. Therefore, the microbial shelf life of the CK group was limited to 20 days, while the ABF group exceeded 30 days. This difference is due to the combined antibacterial effects of WPEO, TiO2, and chitosan in the ABF. The active components in the essential oil, such as terpenes and phenylpropanoids, disrupt bacterial cell membranes, increasing permeability and causing leakage [54,55]. Chitosan also modifies cell membrane permeability, resulting in the leakage of electrolytes and proteins, leading to cell death [56]. TiO2 generates reactive oxygen species through photocatalysis, further compromising cell membrane integrity and ultimately inactivating microorganisms [57,58]. This synergistic effect significantly inhibited microbial growth on the meat surface, delaying the increase in TVC. Ghafari et al. [59] reported that composite films formulated with chitosan (NC), parsley essential oil (ECE), and Iranian tragacanth gum (ITG) exerted strong antibacterial effects against Escherichia coli, Pseudomonas aeruginosa, Staphylococcus aureus, and Bacillus spp. in mutton during refrigerated storage at 4 °C. In a related study, Khodanazary et al. [60] demonstrated that an active nanocomposite film incorporating TiO2, basil essential oil, and polydextrose effectively inhibited the proliferation of several spoilage microorganisms in rainbow trout fillets.
Figure 4. Effects of ABF on the quality characteristics of mutton during super-chilling storage. (A) TVC; (B) TVB-N; (C) TBARS; (D) pH. CK: Control group; CS/WPI/TiO2 + WPEO: Chitosan/Whey protein isolate/TiO2 + White pepper essential oil. Different uppercase letters indicate significant differences (p < 0.05) at the same storage time. Different lowercase letters indicate significant differences (p < 0.05) within the same treatment.

3.2.2. Physicochemical Analysis

TVB-N is commonly defined as volatile nitrogenous bases generated by microbial degradation during storage and is extensively applied as a freshness indicator in meat systems [52,61]. As shown in Figure 4B, the initial TVB-N value was 7.60 mg/100 g. Over time, the TVB-N content progressively rose. However, TVB-N in the ABF group remained significantly lower than in the CK group throughout storage (p < 0.05). Meat is generally regarded as fresh when the TVB-N value does not exceed 15 mg/100 g [62]. At day 25, the TVB-N content of the CK samples rose to 15.17 mg/100 g, exceeding the acceptable limit for fresh meat. In contrast, the TVB-N value of the ABF group remained below 15 mg/100 g at the end of storage, with a value of 14.66 mg/100 g, indicating that ABF effectively inhibited protein degradation and the accumulation of volatile amines. This inhibitory effect can be attributed to the ABF’s strong antimicrobial activity, which aligns with the results observed for TVC. Xu et al. [63] found that pork wrapped in chitosan/polycaprolactone films containing cinnamon essential oil exhibited a 38.78% decrease in TVB-N content after 12 days of refrigerated storage. Similarly, Haghighatpanah et al. [64] reported that TVB-N values significantly decreased when beef samples stored at 4 °C were treated with green bean protein/prulan polysaccharide composite films containing marjoram oil.
Lipid oxidation, caused by the accumulation of lipid peroxides and their secondary metabolites [59], can be monitored by measuring the TBARS. Fresh mutton exhibited a low TBARS value of 0.02 mg/kg, as presented in Figure 4C. Throughout storage, TBARS values increased in both groups; however, samples treated with ABF consistently exhibited lower levels, with significant differences compared with the CK group observed from day 4 onward (p < 0.05). By day 30, TBARS levels in the CK samples increased to 0.52 mg/kg, whereas those in the ABF-treated samples reached only 0.41 mg/kg, indicating that ABF effectively delayed lipid oxidation in mutton. This finding is consistent with previous reports showing that chitosan films incorporating cinnamon essential oil delayed lipid oxidation in chicken [65]. The antioxidant effect of the ABF membrane can be attributed to two main factors. First, WPEO contains antioxidant constituents (e.g., phenolic compounds and terpenes) that can scavenge free radicals, chelate metals, quench singlet oxygen, or donate hydrogen [66,67]. Second, the combined antimicrobial effects of WPEO and TiO2 inhibit microbial growth, which may indirectly reduce lipid oxidation associated with microbial metabolism [31].
The pH value is a crucial indicator for assessing meat freshness [68]. The initial pH was approximately 5.66 (Figure 4D), comparable to values reported previously by Jooyandeh et al. [69]. Throughout storage, the pH of all samples decreased initially and then increased. On day 4, pH in the CK and ABF groups decreased to 5.54 and 5.43, respectively, mainly due to lactic acid accumulation during post-slaughter glycolysis. As storage progressed, the accumulation of volatile basic compounds (e.g., amines and ammonia) produced by microbial metabolism and endogenous enzymatic activity led to a gradual rise in pH [70]. By the end of storage, pH values reached 6.11 in the CK samples, whereas a lower value of 5.86 was observed in the ABF-treated samples. Throughout the storage period, the ABF group consistently exhibited lower pH than the CK group (p < 0.05). This effect is attributed to the functional components of ABF, including chitosan, WPEO, and TiO2, which suppressed the generation of basic nitrogenous compounds associated with microbial activity, thereby retarding pH elevation. Zhu et al. [52] reported that pork packaged with composite films containing 2.5% oregano oil exhibited a lower pH compared with the control group. Similarly, Zhang et al. [3] observed a lower pH in beef treated with agar/sodium alginate films enriched with ginger essential oil relative to the control.

3.2.3. Sensory Evaluation

Sensory evaluation reflects overall quality of a product and its acceptability to consumers, serving as an indicator of the meat’s freshness. Figure 5 presents the appearance of both groups at different storage times. As shown in Figure 5, with increasing storage time, the color of the samples gradually shifted from bright pink to light pink and eventually to brown. This change occurs because, under oxygen exposure and other environmental factors, ferrous iron (Fe2+) in myoglobin is oxidized to ferric iron (Fe3+), leading to metmyoglobin formation. Relative to the CK group, the ABF film more effectively maintained the color of the mutton. Particularly, after 7 days of storage, the fading of pink color was slower in the ABF group, and no browning occurred at the end of storage.
Figure 5. Visual appearance of mutton samples during super-chilling storage. CK: Control group; CS/WPI/TiO2 + WPEO: Chitosan/Whey protein isolate/TiO2 + White pepper essential oil.
Figure 6 shows the sensory scores for both sample groups. The results show that all sensory attributes, including color, odor, texture, and overall acceptability, declined progressively as storage time increased. Within the initial 4 days, sensory scores did not differ significantly between the two treatments (p > 0.05), with all values remaining above 8, reflecting high consumer acceptability. Starting from day 7, the ABF group showed significantly greater sensory scores than the CK group (p < 0.05). By day 25, all sensory scores in the CK group dropped below 4, indicating that the mutton had become unacceptable to consumers. In contrast, sensory scores in the ABF group remained above 5 on 30 days of storage, demonstrating effective preservation of sensory quality. The protective effect of ABF on sensory characteristics can be mainly attributed to the ability of WPEO and TiO2 to suppress microbial proliferation and lipid oxidation, thereby delaying the development of browning and off-flavors in mutton. Additionally, the antimicrobial and antioxidant properties of WPEO in the ABF played an essential role. Similar findings have been reported for composite films containing TiO2 and thyme essential oil used in rainbow trout fillet preservation [60] and for nanocomposite films incorporating 2% cumin oil and 1% TiO2 applied to beef [31].
Figure 6. Changes in sensory quality of mutton during super-chilling storage. (A) Color; (B) odor; (C) texture; (D) overall acceptability. CK: Control group; CS/WPI/TiO2 + WPEO: Chitosan/Whey protein isolate/TiO2 + White pepper essential oil.

4. Conclusions

This study demonstrated that under super-chilling conditions (−1.7 ± 0.2 °C), the ABF containing TiO2 and WPEO exerted a positive influence on the structural and functional properties of MPs, and enhanced mutton quality and shelf life. At the protein level, ABF effectively inhibited protein oxidation, as evidenced by slowing down the reduction in total sulfhydryl content and protein solubility, and retarding the increase in carbonyl content and surface hydrophobicity. Moreover, the ABF contributed to the structural integrity of MPs, as evidenced by a higher proportion of α-helices, reduced random-coil content, a smaller decline in intrinsic fluorescence intensity, and a more compact gel microstructure. In terms of mutton quality, ABF demonstrated excellent antioxidant and antimicrobial activities significantly delaying the rise in TVC, TBARS, and TVB-N, thus extending the shelf life by over 10 days compared with the CK group. Therefore, the ABF effectively protects protein structure and delays meat quality deterioration during super-chilling storage, demonstrating its potential for mutton preservation.

Author Contributions

Conceptualization, Y.Z. and R.C.; methodology, R.C. and Y.D.; software, Y.D.; validation, R.C.; formal analysis, R.C.; investigation, R.C.; resources, Y.Z.; data curation, R.C.; writing—original draft preparation, R.C.; writing—review and editing, Y.Z. and R.C.; visualization, R.C. and Y.D.; supervision, Y.Z.; project administration, Y.Z.; funding acquisition, Y.Z. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the Key Research and Development Program of Shanxi Province (202302140601017) and the Construction Project of the Functional Food Industry Technology System of Shanxi Province (2024CYJSTX10-03).

Institutional Review Board Statement

The sensory evaluation conducted in this study adheres to standard food safety protocols and does not involve any intervention with human subjects. According to the Notice on the Administration of Ethical Review of Life Science and Medical Research Involving Humans (National Health Commission of China, 2023), ethical approval is not required for studies that focus solely on sensory evaluation of food products. Therefore, no ethics committee approval was needed for this study.

Data Availability Statement

The original contributions presented in this study are included in the article. Further inquiries can be directed to the corresponding author.

Conflicts of Interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

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