Effects and Mechanisms of Ultrasonic and Papain Treatment on Texture Properties of the Abalone (Haliotis discus hannai) Meat

Abstract

Haliotis discus hannai meat has a firm texture that makes it difficult to chew and swallow, so tenderizing is necessary. Ultrasonic treatment and papain enzyme processing are used to reduce the hardness of abalone meat. This study tests physicochemical indicators and protein changes to assess meat quality and protein alterations. The maximum reduction in hardness of raw meat reached 60.58%, while heat-treated abalone meat achieved 61.13%, with free water and bound water converting to immobile water. The L* value of the meat decreased, while the a* and b* values increased. In raw meat, the content of TCA-soluble peptides increased with increasing treatment intensity. However, in heat-treated meat, this peptide content decreased with rising temperature. Muscle fiber filament breaks and pore numbers increased. The BPB binding content showed a negative correlation with the percentage of α-helix. Total sulfhydryl and free amino groups in raw meat decrease with increasing treatment intensity; both parameters in cooked meat decrease with rising temperature. Changes in tertiary protein structure cause alterations in fluorescence intensity, with secondary structure shifting from α-helix to β-sheet conformation. The results suggested that ultrasonic and papain treatments can induce structural alterations in proteins. This leads to protein cleavage and depolymerization, collectively resulting in softening of abalone meat texture and redistribution of internal moisture. These processes result in softening of abalone meat and redistribution of internal moisture. This study provides a theoretical basis for developing abalone tenderizers.

1. Introduction

Haliotis discus hannai is favored by consumers both domestically and internationally, boasting broad prospects for market development. It is widely distributed along the coasts of China, Japan, and the Korean Peninsula. The edible portion of abalone meat contains 30–50% myosin protein (MP) [1]. However, the firm and tough texture of abalone meat limits its consumption among specific consumer groups, making research on abalone meat tenderization imperative. As a class of natural proteins, enzymes are characterized by non-toxicity, high reaction efficiency, and cost-effectiveness. Furthermore, previous studies have confirmed that enzymes play a crucial role in meat tenderization, with plant proteases being recognized as a green approach to improving meat texture [2]; for example, Ficin, which remains active in acidic environments [3], Ginger protease, with specificity for collagen [4], and papain, used in past studies. The ability of papain to improve beef tenderness has been demonstrated, attributed to the degradation of myofibrillar proteins and disruption of muscle fiber structure [5,6]. Heba H.S. Abdel-Naeem et al. [7] have used papain and ginger extracts to treat camel meat to destroy its muscle fiber and connective tissues, which resulted in a tenderizing effect on camel meat. The widespread use of food-grade enzymes also proves that the safety of enzymes is unquestionable.

Ultrasound technology has many advantages, such as being green, safe, inexpensive, and efficient, and can contribute to meat tenderization through the cavitation effect of ultrasound. In addition, previous studies have investigated the effects of ultrasound and enzymes in the tenderization of aquatic products, characterizing the effects on the flavor and quality of the raw material. Ultrasound technology may be a useful means to achieve enzyme diffusion in meat [8]. Du et al. [9] had treated chicken meat using 500 W ultrasound power for 30 min and reduced the shear force by 27.10% compared to the control. Xiong et al. [10] also employed ultrasonic treatment on chicken meat to investigate its effect on tenderness. Following combined ultrasonic and sodium bicarbonate treatment, the shear force of chicken meat decreased while the myofibrillar rupture index increased. Buccinum undatum is a marine mollusk belonging to the Gastropoda class. Treatment with 200 W, 45 kHz ultrasound for 9.6 min significantly reduces the shear force of abalone meat and enhances its tenderness [11]. Haliotis discus hannai, also a marine mollusk within the Gastropoda class, may similarly experience reduced hardness when subjected to ultrasonic processing. The application of ultrasonic technology in meat tenderization holds substantial promise, particularly within the domain of meat processing. Specifically, the cleavage of myofibrillar proteins, degradation of collagen, and reduction in the diameter of myofibrillar bundles have been identified as plausible mechanisms underlying ultrasound-induced meat tenderization [12].

As a major protein component in the edible part of abalone, myofibrillar proteins represent a critical focus for investigating the mechanisms underlying abalone meat tenderization induced by these two treatments. Previous studies have been conducted to reveal the mechanism of meat tenderization through the observation of changes in myofibrillar proteins. Zhang et al. [13] tenderized yak meat by lactic acid-ultrasound-papain combination; the denaturation of myofibrillar proteins and alteration of spatial structure were observed and considered to be an important factor in the improvement of the quality of yak meat. Shi et al. [14] improved the tenderness of chicken breast meat by using potassium alginate and ultrasonic technology. Ultrasonic treatment increased intermuscular spaces, facilitating potassium alginate penetration. The combined effect formed a surface barrier that prevented tissue shrinkage and dehydration, thereby enhancing meat tenderness.

In this study, the mechanisms of ultrasound and enzymes on the tenderization of aquatic products were investigated through systematic experiments. It also elucidated the mechanism of meat tenderness change from the point of view of myofibrillar protein change and explored the texture and moisture change of abalone meat under different heating temperatures based on the optimal group of texture improvement. It is hoped that the mechanisms and research findings revealed by this work will provide insights for the subsequent optimization and upgrading of abalone meat tenderization technology based on ultrasonic-enzyme synergy. It will offer theoretical foundations and scientific references for studies on process parameter control and efficacy enhancement of this technology.

2. Materials and Methods

2.1. Materials

Fresh abalone were purchased from local aquatic supermarkets in Pudong New Area, Shanghai, and transported to the laboratory for dissecting and killing within 0.5 h under ice.

Trichloroacetic acid (TCA, standard for AR), 5,5′-dithio (2-nitrobenzoic acid) (DTNB, standard for AR), Bromophenol blue (standard for AR), Potassium bromide (standard for SP), and o-Phthalaldehyde (OPA, standard for GC) were purchased from Shanghai Macklin Biochemical Co., Ltd. (Shanghai, China); papain (food-grade; ≥800 u/mg) was purchased from Shanghai Xinyu Bio-Tech. Ltd. (Shanghai, China).

2.2. Methods

2.2.1. Pre-Processing

Based on the number of groups and taking ten abalone from each group, the weight of each abalone is about 70~80 g. The abalone meat was completely submerged in phosphate (20 mM Na₂HPO₄-NaH₂PO₄ pH 7.0) solution (w/w 1:4), recorded as CK. The number of killed abalone was divided into two groups, both completely submerged in phosphate buffer solution, then transferred to an ultrasonic cell disruptor for sonication. The ultrasonic probe was submerged 2 cm below the liquid surface. Ultrasonic treatment was performed using an ultrasonic cell disruptor (SX-2008, SHANG HAI SX INSTRUMENT CO., LTD., Shanghai, China) equipped with an ultrasonic horn. To control temperature, the ultrasonic treatment process was conducted in a cold bath. Ultrasound was performed at 1000 W power and 20 kHz frequency for 20 and 30 min, designated as U2 and U3, respectively. Concentrations of 3% and 5% (w/v) papain were configured using the phosphate solution, according to the enzyme solution and abalone meat 3:1 (w/w), and completely submerged with magnetic stirring for 6 h, noted as A3 and A5. A3 and A5 were wiped and submerged in phosphate solution, respectively, and transferred to a cell ultrasonic breaker, where the same ultrasound conditions were treated, noted as AU32, AU33, AU52, and AU53. Reversing the order of ultrasonication and enzyme submerging, the processed abalone meat was noted as UA32, UA33, UA52, and UA53. Abalone in groups CK, AU53, and UA53 were selected and immersed in distilled water according to 5:1 (w/w) and heated to the center temperatures of 50, 60, 70, and 80 °C, respectively, for 10 min. The samples were recorded as B\U\R-5\6\7\8.

2.2.2. Determination of TPA

The texture analysis (TPA) of meat samples was measured using the method described by Fang et al. [15]. The texture testing instrument selected is the TA. XT plus, Stable Micro System, Godalming, UK. Considering the species differences, the method was then slightly modified. Abalone meat was cut into small pieces of 1*1*1 cm and stabilized at room temperature for 1 h before measurement using a P/50 probe with 2 g trigger force, 10 mm test distance, 1 mm/s pre-test speed, 1 mm/s test speed, and 30% compression ratio. Each sample was tested 3 to 5 times.

2.2.3. Determination of Centrifugal Loss

The method was slightly modified according to Yang et al. [16]. A total of 1 g abalone meat sample was weighed, wrapped in filter paper, and centrifuged at 10,000 rpm at 4 °C for 10 min. The weight after centrifugation was weighed and calculated by the following formula:

$$\mathrm{C}\mathrm{e}\mathrm{n}\mathrm{t}\mathrm{r}\mathrm{i}\mathrm{f}\mathrm{u}\mathrm{g}\mathrm{a}\mathrm{l} \mathrm{l}\mathrm{o}\mathrm{s}\mathrm{s} (\%)={\displaystyle \frac{\left(\mathrm{W}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t} \mathrm{b}\mathrm{e}\mathrm{f}\mathrm{o}\mathrm{r}\mathrm{e} \mathrm{c}\mathrm{e}\mathrm{n}\mathrm{t}\mathrm{r}\mathrm{i}\mathrm{f}\mathrm{u}\mathrm{g}\mathrm{a}\mathrm{l}\left(\mathrm{g}\right)-\mathrm{W}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t} \mathrm{a}\mathrm{f}\mathrm{t}\mathrm{e}\mathrm{r} \mathrm{c}\mathrm{e}\mathrm{n}\mathrm{t}\mathrm{r}\mathrm{i}\mathrm{f}\mathrm{u}\mathrm{g}\mathrm{a}\mathrm{l}\left(\mathrm{g}\right)\right)}{\left(\mathrm{W}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t} \mathrm{b}\mathrm{e}\mathrm{f}\mathrm{o}\mathrm{r}\mathrm{e} \mathrm{c}\mathrm{e}\mathrm{n}\mathrm{t}\mathrm{r}\mathrm{i}\mathrm{f}\mathrm{u}\mathrm{g}\mathrm{a}\mathrm{l}\left(\mathrm{g}\right)\right)}}$$

2.2.4. Water Distribution

The method was slightly modified according to Zhou et al. [17]. The parameters were recalibrated to take sample differences into account. The water distribution in the sample was determined by measuring the transverse relaxation time (T₂). The measurements of the transverse relaxation time were performed on a MesoMR23-060H-I pulse NMR analyzer (Shanghai Niumai Electronic Technology Co., Shanghai, China). The gel samples were placed in the NMR tube at 32 °C with a proton resonance frequency of 21 MHz, an echo time of 2 ms, and an echo number of 1000, and the data were inverted iteratively to make a graph. After wrapping the sample with plastic film, it was put into the MRI tube for imaging considering the sample differences, so the parameters were corrected. The pseudo-color map was processed by the image software of Shanghai Newmax Co. (Niumag NMR Image Processing Software V3.0, Shanghai Niumai Electronic Technology Co., Shanghai, China).

2.2.5. Determination of Cooking Loss

The method was slightly modified according to Eddin a et al. [18]. A 1 g sample of abalone meat was weighed and heated in water at 90 °C for 30 min in isolation. The surface was dried and weighed, and the cooking loss was calculated by the following formula:

$$\mathrm{C}\mathrm{o}\mathrm{o}\mathrm{k}\mathrm{i}\mathrm{n}\mathrm{g} \mathrm{l}\mathrm{o}\mathrm{s}\mathrm{s} (\%)={\displaystyle \frac{(\mathrm{W}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t} \mathrm{b}\mathrm{e}\mathrm{f}\mathrm{o}\mathrm{r}\mathrm{e} \mathrm{h}\mathrm{e}\mathrm{a}\mathrm{t}\mathrm{i}\mathrm{n}\mathrm{g}\left(\mathrm{g}\right)-\mathrm{W}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t} \mathrm{a}\mathrm{f}\mathrm{t}\mathrm{e}\mathrm{r} \mathrm{h}\mathrm{e}\mathrm{a}\mathrm{t}\mathrm{i}\mathrm{n}\mathrm{g} (\mathrm{g}\left)\right)}{(\mathrm{W}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t} \mathrm{b}\mathrm{e}\mathrm{f}\mathrm{o}\mathrm{r}\mathrm{e} \mathrm{h}\mathrm{e}\mathrm{a}\mathrm{t}\mathrm{i}\mathrm{n}\mathrm{g} (\mathrm{g}\left)\right)}}$$

2.2.6. Determination of Color

Testing of the color change of abalone meat was performed using a colorimeter (CR-40, Konica Minolta Sensing Inc., Tokyo, Japan). Several samples of abalone meat were taken and tested after wiping the surface moisture. The L*, a* and b* values of abalone meat were recorded. Each sample was tested 3 to 5 times.

2.2.7. Determination of pH

A total of 2 g of abalone meat were placed in distilled water, homogenized using a homogenizer, and then centrifuged to collect the supernatant. A pH meter (FE20, Mettler Toledo Co., Shanghai, China) was used to test the pH value of the supernatant.

2.2.8. Microstructure

Freshly processed abalone meat samples were cut into small pieces and fixed in 20% glutaraldehyde solution for 24 h. The samples were dehydrated using gradient concentration of ethanol solution and sprayed with gold for 1 min (10 mA); then, the morphology of the samples was observed and recorded under the Hitachi SU5000 thermal field emission scanning electron microscope(HITACHI., Tokyo, Japan). The magnification of this electron microscope is 400×. Freshly processed abalone meat samples were cut into small pieces and stored in formalin solution, fixed in paraffin wax, and sliced. The slices were processed in constant dye pretreatment solution for 1 min, then dehydrated with 95% ethanol for 1 min, stained with eosin staining solution for 15 s, dehydrated and processed after staining, and then sealed and examined microscopically under a light microscope MS500W (Shanghai Metz Precision Instruments Co., Ltd., Shanghai, China). The magnification of this optical microscope is 200×.

2.2.9. Determination of TCA-Soluble Peptides

The method was slightly modified according to Zhang et al. [19]. A total of 2 g of fresh abalone meat sample was placed in 18 mL of 5% (w/v) Trichloroacetic acid (TCA) solution, churned using a homogenizer, equilibrated in an ice bath for 30 min, and centrifuged at 10,000 rpm for 15 min at 4 °C. Using the BCA method, absorbance was measured at 540 nm using a spectrophotometer (SP-756P, Shanghai Spectrometer Instruments Co., Ltd., Shanghai, China).

2.2.10. Determination of SDS-PAGE

A total of 3 g of abalone meat was taken, placed in 5% (w/v) SDS solution, homogenized and centrifuged to take the supernatant, and heated in a boiling water bath for 30 min. The sample buffer was mixed and then heated in a boiling water bath for 10 min. Tris-Gly electrophoresis precast gels and Marker (6.5~270 kDa) were purchased from Beyotime, and the volume of Marker and specimen samples was 8 μL. The initial voltage was 80 V. After 30 min, the voltage was changed to 120 V, and the sample was continued for 90 min. at 80 V; after 30 min, the voltage was changed to 120 V and continued for 90 min. The gels were stained with Caumas Brilliant Blue R250, and after 1~2 h, the dye was removed with a decolorizing solution and observed by overnight immersion in distilled water.

2.2.11. Myofibrillar Protein Extraction

The method was slightly modified according to Wang et al. [20]. The abalone meat was equally divided in 5 times volume of a low salt washing solution (pH 7.0) consisting of 0.1 M NaCl, 20 mM PBS, 2 mM MgCl₂, and 1 mM EDTA, and 10,000 rpm homogenization for 30 s was repeated three times, with a 20 s pause in between. Centrifugation at 10,000 rpm, 4 °C was performed for 15 min and then repeated on the precipitate four times. The precipitate was taken in 5 times volume of high salt extract (pH 7.0), which consisted of 0.6 M NaCl, 20 mM PBS, 2 mM MgCl₂, and 1 mM EDTA, homogenized under the same conditions and repeated. The supernatant was taken after centrifugation under the same conditions after standing for 1 h. A 10 times volume of pre-cooled deionized water was washed and centrifuged to take the precipitate as crude myofibrillar protein. The protein was stored at 4 °C and used up within 48 h.

2.2.12. Determination of BPB Binding Content

The method was slightly modified according to Wang et al. [21]. The extracted myofibrillar protein solution was diluted to 1 mg/mL. A total of 80 μL of Bromophenol Blue (BPB) solution at a concentration of 1 mg/mL was aspirated, added dropwise to 2 mL of protein lysate or buffer, shaken for 10 min, and centrifuged at 10,000 rpm for 15 min at 4 °C. The supernatant was taken and tested for absorbance at 595 nm. The hydrophobicity was calculated as the amount of BPB bound by the following formula:

$$\mathrm{B}\mathrm{P}\mathrm{B} \mathrm{b}\mathrm{i}\mathrm{n}\mathrm{g}\mathrm{d}\mathrm{i}\mathrm{n}\mathrm{g} \mathrm{c}\mathrm{o}\mathrm{n}\mathrm{t}\mathrm{e}\mathrm{n}\mathrm{t}\left(\mathrm{\mu }\mathrm{g}\right)={\displaystyle \frac{80 \mathrm{\mu }\mathrm{g}\ast \left({\mathrm{O}\mathrm{D}}_{\left(\mathrm{c}\mathrm{o}\mathrm{n}\mathrm{t}\mathrm{r}\mathrm{o}\mathrm{l}\right)}-{\mathrm{O}\mathrm{D}}_{\left(\mathrm{e}\mathrm{x}\mathrm{p}\mathrm{e}\mathrm{r}\mathrm{i}\mathrm{e}\mathrm{n}\mathrm{m}\mathrm{e}\mathrm{n}\mathrm{t}\right)}\right)}{\left({\mathrm{O}\mathrm{D}}_{\left(\mathrm{c}\mathrm{o}\mathrm{n}\mathrm{t}\mathrm{r}\mathrm{o}\mathrm{l}\right)}\right)}}$$

2.2.13. Determination of Fluorescence Spectrum

The method was slightly modified according to Guo et al. [22]. Fluorescence spectroscopy scanning was performed using a fluorescence spectrometer (Hitachi F-7100 3D Fluorescence Spectrometer, HITACHI., Tokyo, Japan). The extracted myofibrillar protein solution was diluted to a protein content of 0.1 mg/mL using 0.6 M NaCl solution and fluorescence scanning solution with excitation and emission slit bandwidths of 10 nm and 5 nm, respectively. The excitation wavelength was 280 nm, and the emission wavelength was 290–450 nm.

2.2.14. Determination of Total Sulfhydryl and Free Amino Groups Content

The method was slightly modified according to Hu et al. [23]. Diluted myofibrillar protein solution was used (0.2 M Tris-HCl, 8 M urea, 3 mM EDTA, 1% SDS, pH 8.0) to dilute the protein concentration to 1 mg/mL. A total of 2 mL of diluted protein solution was mixed with 50 μL of Ellman’s reagent (10 mM DTNB, 0.2 M Tris-HCl, pH 8.0) at 40 °C, protected from light. After holding for 25 min, the absorbance was tested at 412 nm by a Spectrophotometer (SP-756P, Shanghai Spectrometer Instruments Co., Ltd., Shanghai, China). The molar extinction coefficient was recorded as 13,600. The total sulfhydryl content was calculated by the following formula:

$$\mathrm{T}\mathrm{o}\mathrm{t}\mathrm{a}\mathrm{l} \mathrm{s}\mathrm{u}\mathrm{l}\mathrm{f}\mathrm{h}\mathrm{y}\mathrm{d}\mathrm{r}\mathrm{y}\mathrm{l} \mathrm{g}\mathrm{r}\mathrm{o}\mathrm{u}\mathrm{p} \mathrm{c}\mathrm{o}\mathrm{n}\mathrm{t}\mathrm{e}\mathrm{n}\mathrm{t} (\mathrm{\mu }\mathrm{m}\mathrm{o}\mathrm{l}/\mathrm{g} \mathrm{p}\mathrm{r}\mathrm{o}\mathrm{t}\mathrm{e}\mathrm{i}\mathrm{n})={\displaystyle \frac{\mathrm{A}\mathrm{b}\mathrm{s}\mathrm{o}\mathrm{r}\mathrm{b}\mathrm{a}\mathrm{n}\mathrm{c}\mathrm{e} \ast \mathrm{D}\mathrm{i}\mathrm{l}\mathrm{u}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n} \mathrm{r}\mathrm{a}\mathrm{t}\mathrm{i}\mathrm{o} \ast \mathrm{100,000}}{(\mathrm{13,600} \ast \mathrm{C}\mathrm{u}\mathrm{v}\mathrm{e}\mathrm{t}\mathrm{t}\mathrm{e} \mathrm{t}\mathrm{h}\mathrm{i}\mathrm{c}\mathrm{k}\mathrm{n}\mathrm{e}\mathrm{s}\mathrm{s} \ast \mathrm{P}\mathrm{r}\mathrm{o}\mathrm{t}\mathrm{e}\mathrm{i}\mathrm{n}\mathrm{c}\mathrm{o}\mathrm{n} \mathrm{c}\mathrm{e}\mathrm{n}\mathrm{t}\mathrm{r}\mathrm{a}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n})}}$$

The method of free amino groups was slightly modified according to Bao et al. [24]. The amount of free amino groups was tested using the OPA method. A total of 160 mg of ortho- Phthaldialdehyde (OPA) was dissolved in 4 mL of anhydrous ethanol. It was transferred to 100 mM borax solution (10% SDS, 176 mg DTT) and fixed with deionized water to a final volume of 200 mL. Protein concentration was diluted to 1 mg/mL and mixed with OPA solution (1:6). The absorbance at 340 nm was measured, and the standard curve was plotted with glycine.

2.2.15. Determination of FT-IR

We employed Fourier Transform Infrared Spectroscopy (FT-IR) to assess changes in protein secondary structure. The secondary structure was modified according to Zhang et al. [25]. The lyophilized protein powder, mixed according to protein powder: potassium bromide 1:100 (w/w) and pressed into tablets, was analyzed by removing the background using an FT-IR infrared spectrometer (model), and transmission spectra were recorded in the wavelength range of 4000–500 cm⁻¹. After preliminary processing of the curves using OMINIC (OMINIC32, Thermo Fisher Scientific Inc., Waltham, MA, USA), the peaks were split and fitted to the 1700–1600 cm⁻¹ peaks using Peakfit v4.12 to calculate the percentage of secondary structure.

2.3. Data Analysis

Data were analyzed using IBM SPSS Statistics 27, ANOVA was used for analysis of variance and significance, and significant differences between treatment groups were compared using Duncan’s method for multiple comparisons, with p < 0.05 indicating significant differences between groups. Each part of the experiment was repeated three times independently. Data were expressed as mean ± standard deviation. Graphs were plotted using Origin 64 software.

3. Results and Discussion

3.1. TPA

The textural properties of the tested samples can simulate the action of the human mouth and teeth on abalone meat, and the analysis of the textural properties can illustrate the effect of the tenderizing method on abalone meat to obtain the best processing technique. Among them, hardness is an important index to measure the effect of tenderization. As shown in Table 1, the addition of enzyme and the increase in concentration decreased the hardness of abalone meat, and the hardness of abalone meat in the A3 and A5 groups decreased by 12.64% and 31.37%, respectively, relative to the CK group. Xu et al. [26] used pineapple protease and papain to tenderize giant squid meat and also found a significant decrease in meat hardness and shear force. They concluded that the protein hydrolysis activity of the enzymes led to the breakage of myofibrillar proteins and disruption of connective tissues, which would result in a decrease in hardness. Ultrasound application and prolonged treatment time gave similar results, with a reduction in abalone meat hardness of 29.87% and 40.60% in the U2 and U3 groups, respectively. Peña-Gonzalez et al. [27]. investigated the effect of ultrasound on the physical characteristics of beef, and it was found that ultrasound significantly reduced the hardness of beef. In addition, ultrasound induces changes in macromolecules such as enzymes, which keep them highly active, and consequently affects the hardness of meat. After the combined effect of enzymes and ultrasound, the hardness of each group was significantly reduced compared to the CK group. The hardness of abalone meat in the AU32, AU33, AU52, and AU53 groups was reduced by 37.93, 54.24, 43.94, and 60.58%, respectively, compared to the CK group. The hardness of abalone meat in the UA32, UA33, UA52, and UA53 groups was reduced by 37.81, 48.29, 46.12, and 54.59%. Except for the UA52 group, the hardness reduction of the other groups was not as effective as that of the AU group. In addition, elasticity was reduced in all experimental groups compared to the control, but the difference was not significant. Adhesion and chewability were significantly reduced in all groups compared to the control, with the highest reductions of 69.50% and 77.03% and the lowest reductions of 53.30% and 63.44% in the AU group, respectively. The UA group had similar effects. Ultrasound and enzymes will disrupt the fibrous structure of the meat; the cavitation effect of ultrasound cuts the meat fiber filaments, and enzymes disrupt the protein structure of the meat. The ultrasound action also activates the proteases inherent in the meat, leading to degradation of the proteins [28]. Bao et al. [29] used different powers of ultrasound to treat beef and found that ultrasound powers above 200 W resulted in a significant reduction in hardness of beef, and they concluded that cytoskeletal changes and denaturation of proteins were the main reasons.

Table 1. Effects of ultrasound and enzyme treatments on textural properties of abalone meat before and after treatment.

	Hardness (g)	Adhesiveness	Springiness	Cohesiveness	Gumminess	Chewiness (g)	Resilience
CK	1841.08 ± 86.14 a	−14.9 ± 7.78 abc	0.92 ± 0.03 b	0.58 ± 0.06 cd	1062.76 ± 97.55 a	981.22 ± 114.92 a	0.4 ± 0.05 a
A3	1608.37 ± 100.01 b	−16.75 ± 3.12 d	0.76 ± 0.01 b	0.49 ± 0.09 de	790.77 ± 201.63 bcd	603.27 ± 157 cd	0.34 ± 0.07 a
A5	1263.6 ± 149.64 c	−9.83 ± 4.46 cd	0.71 ± 0.01 a	0.48 ± 0.07 de	593.73 ± 108.48 de	431.21 ± 98.24 def	0.33 ± 0.05 a
U2	1291.09 ± 104.57 c	−5.83 ± 1.47 ab	0.65 ± 0.03 a	0.62 ± 0.03 c	798.16 ± 94.17 bcd	521.75 ± 82.83 cde	0.4 ± 0.05 a
U3	1093.65 ± 9.80 de	−12.11 ± 0.42 bcd	0.61 ± 0.02 a	0.75 ± 0.10 b	817.6 ± 105.69 bc	499.9 ± 70.26 cde	0.38 ± 0.08 a
AU32	1142.82 ± 73.61 cd	−7.89 ± 4.29 abc	0.72 ± 0.05 a	0.43 ± 0.02 e	496.05 ± 58.40 ef	358.73 ± 48.59 efg	0.31 ± 0.03 a
AU33	842.52 ± 33.45 fg	−5.51 ± 8.68 ab	0.65 ± 0.01 a	0.41 ± 0.11 e	417.06 ± 53.95 ef	265.31 ± 30.94 fg	0.5 ± 0.31 a
AU52	1032.07 ± 72.20 de	−7.73 ± 30 abc	0.7 ± 0.15 a	0.46 ± 0.10 de	482.11 ± 135.14 ef	356.05 ± 168.39 efg	0.3 ± 0.04 a
AU53	725.67 ± 49.59 g	−4.07 ± 0.63 a	0.66 ± 0.08 a	0.44 ± 0.00 e	323.92 ± 22.95 f	225.32 ± 5.54 g	0.3 ± 0.01 a
UA32	1145.39 ± 45.98 cd	−5.38 ± 2.37 ab	0.89 ± 0.04 a	0.86 ± 0.02 ab	980.93 ± 39.90 e	875.53 ± 38.61 ab	0.39 ± 0.03 a
UA33	952.28 ± 28.31 ef	−6.29 ± 1.68 ab	0.95 ± 0.01 a	0.88 ± 0.03 a	840.98 ± 55.11 bc	795.55 ± 55.82 ab	0.43 ± 0.01 a
UA52	992.22 ± 13.30 def	−4.11 ± 0.78 a	0.94 ± 0.03 a	0.87 ± 0.03 ab	860.11 ± 40.49 bc	807.95 ± 59.91 ab	0.41 ± 0.04 a
UA53	836.97 ± 5.72 fg	−3.75 ± 0.87 a	0.93 ± 0.020 a	0.89 ± 0.02 a	741.7 ± 17.09 cd	691.85 ± 29.73 bc	0.42 ± 0.00 a

The same superscript capital letters (a–g) in a column indicate no significant differences in the column (p > 0.05).

Table 2 shows the changes in the textural properties of heat-treated abalone meat. After heat treatment, the hardness of abalone meat exhibited a gradual decrease with increasing temperature. The lowest hardness of untreated abalone meat was 610.67 ± 15.52 N in the B8 group, and the lowest hardness of treated abalone meat was 237.41 ± 31.52 N in the R8 group, both of which showed the lowest hardness at 80 °C. Compared to B8, the hardness of U8 and R8 decreased by 53.26% and 61.13%, respectively. In addition, at the same heating temperature, the hardness of the treated groups was lower than that of the untreated group, and there was a difference in hardness change between the U and R groups. During heating, muscle tissue undergoes structural changes, including lateral or longitudinal contraction of myofibers, gelation of protein aggregates, shortening and dissolution of connective tissue, and dissolution of collagen. These changes collectively contribute to affect tenderness [30]. Heating-induced slow contraction of muscle fiber filaments facilitates moisture retention, thereby contributing to a reduction in meat hardness. High-intensity ultrasound pretreatment induces fiber contraction, which further stabilizes the fibrous filaments during subsequent heating, thereby facilitating moisture retention and reducing firmness simultaneously. At the protein level, protein aggregation—evidenced by an increase in molecular weight—was also observed in SDS-PAGE experiments, further confirming the role of high-intensity ultrasound pretreatment in regulating water-holding capacity and hardness. However, the reasons for the changes in hardness between treatment groups still need to be verified in subsequent experiments.

Table 2. Effect of heat treatment on textural properties of cooked abalone meat after heat treatment.

	Hardness (g)	Adhesiveness	Springiness	Cohesiveness	Gumminess	Chewiness (g)	Resilience
B5	1191.73 ± 15.16 a	−19.41 ± 11.14 ab	1.02 ± 0.08 a	0.9 ± 0.04 abc	1070.38 ± 57.64 a	1097.5 ± 130.56 a	0.41 ± 0.02 a
B6	807.33 ± 16.57 c	−0.74 ± 0.17 a	0.99 ± 0.01 ab	0.92 ± 0.02 ab	741.01 ± 29.96 c	732.22 ± 34.75 bc	0.55 ± 0.02 b
B7	622.3 ± 4.34 de	−1.89 ± 0.26 a	1 ± 0.01 ab	0.94 ± 0 a	582.4 ± 6.95 d	579.92 ± 9 d	0.6 ± 0.01 a
B8	610.67 ± 15.52 e	−0.88 ± 0.41 a	0.99 ± 0.01 ab	0.92 ± 0.03 ab	561.77 ± 19.63 de	557.02 ± 26.56 de	0.57 ± 0.03 ab
U5	947.63 ± 57.19 b	−37.16 ± 16.13 bc	0.94 ± 0.05 bc	0.89 ± 0.05 abcd	845.09 ± 89 b	800.75 ± 121.33 b	0.45 ± 0.03 d
U6	555.34 ± 26.22 f	−1.13 ± 0.18 a	0.99 ± 0.01 ab	0.92 ± 0.01 ab	511.63 ± 27.58 e	506.84 ± 32.99 de	0.5 ± 0 c
U7	477.79 ± 8.44 f	−52.23 ± 3.09 c	0.99 ± 0.01 ab	0.87 ± 0.03 abcd	399.5 ± 38.25 f	393.5 ± 36 fg	0.37 ± 0.02 ef
U8	285.41 ± 10.93 h	−10.68 ± 0.88 ab	0.9 ± 0.01 c	0.85 ± 0.03 cd	243.81 ± 16.17 g	220.59 ± 16.94 h	0.35 ± 0.02 f
R5	808.03 ± 6.8 c	−14.2 ± 1.42 d	0.94 ± 0.03 bc	0.88 ± 0.04 abcd	714.44 ± 30.14 c	670.77 ± 45.17 c	0.38 ± 0.04 ef
R6	649.33 ± 20.14 d	−19.93 ± 3.06 e	0.88 ± 0.05 c	0.83 ± 0.09 d	536.19 ± 43.72 de	473.8 ± 59.35 ef	0.37 ± 0.04 f
R7	446.61 ± 29.87 f	−15.98 ± 1.58 d	0.9 ± 0.04 c	0.87 ± 0.03 bcd	387.66 ± 34.52 f	347.5 ± 34.8 g	0.29 ± 0.03 g
R8	237.41 ± 31.52 i	−15.16 ± 3.14 d	0.9 ± 0.07 c	0.84 ± 0.02 cd	199.11 ± 28.95 g	179.08 ± 30.19 i	0.26 ± 0.04 g

The same superscript capital letters (a–i) in a column indicate no significant differences in the column (p > 0.05).

3.2. Centrifugal Loss and Water Distribution

Centrifugal loss reflects a meat product’s ability to hold its original moisture when exposed to external forces. The structure of protein molecules and the integrity of muscle fibers are key factors that affect changes in water-holding capacity. Thus, depolymerization and cleavage of protein molecules will necessarily weaken water-holding capacity, leading to an increase in centrifugal loss. As shown in Figure 1B, in the AU and UA groups, centrifugal loss was reduced by a maximum of 3.04% and a minimum of 2.33% compared to the CK group. Centrifugal loss was lower in the AU and U groups compared with the CK group, but higher in the A group than in the CK group. In cooked meat, as shown in Figure 2B, centrifugal loss gradually decreased with increasing heating temperature in groups U and B, with no significant change in group R. In the U and B groups, centrifugal loss dropped from 8.27% in B5 to 4.57% in B8, and from 5.97% in U5 to 3.83% in U8. In both raw and cooked samples, centrifugal loss was lower in abalone meat treated with ultrasound than in the untreated group. However, the centrifugal loss of raw meat treated solely with enzymes was higher than that of the CK group. This phenomenon may be due to the cavitation effect of ultrasound, which destroys myogenic fibers and alters the protein structure of meat to form fine ordered spaces, which in turn retain more water [31]. Enzymes cause proteolysis, which breaks down the ordered structure and thus impairs moisture retention.

Figure 1. Changes in cooking loss (A), centrifugal loss (B), TCA-soluble peptides (C), total sulfhydryl content (D), free amino groups content (E), BPB binding content (F), L* (G), a* (H), b* (I), pH (J), and moisture distribution (K,L) of raw abalone meat prior to heat treatment. Note: different lowercase letters indicate significant differences (p < 0.05).

Figure 2. Changes in cooking loss (A), centrifugal loss (B), TCA-soluble peptide (C), total sulfhydryl content (D), free amino groups content (E), BPB binding content (F), L* (G), a* (H), b* (I), pH (J), and moisture distribution (K,L) of cooked abalone meat after heating treatment. Note: different lowercase letters indicate significant differences (p < 0.05).

LF-NMR allows the determination of water distribution in foods. The NMR curves of each group are shown in Figure 1 and Figure 2K. T2a and T2b both represent bound water in meat, with relaxation times ranging from 1 to 10 ms; T22 represents fixed water, with a relaxation time of about 100 ms; and T23 represents free water, with a relaxation time of about 1000 ms [32]. Figure 1 and Figure 2L demonstrate the percentage of various types of water in raw and cooked meat samples, respectively. As shown in Figure 1L, the AU53 group had the highest percentage of immobile water at 95.73%. Immobile water refers to the water between the protein networks of myofibrils. It made up the largest proportion in the meat. The CK group showed an immobile water percentage of 95.23%. Enzyme treatment reduced this percentage to 95.21% and 94.84%. Protease cleaves myofibrillar protein molecules and destroys the original network structure, resulting in the loss or transfer of water [33]. Compared with the CK group, the proportion of bound water in the A3 group went up from 3.77% to 3.89%, while the free water in the A5 group increased from 1.00% to 1.21%. This suggests that water shifts from immobile water to both free water and bound water. However, in the AU group, higher enzyme concentration and longer ultrasound time caused a reduction in free water. For example, the percentages of free water and bound water in AU32 were 1.27% and 3.81%, respectively, versus 1.08% and 3.43% in AU52. Meanwhile, the immobile water percentage was 94.91% in AU32, compared with 95.50% in AU52. The same trend was observed in AU33 and AU53. The UA group showed similar results to the AU group. Thus, the combined treatment caused water to shift from bound and free water to immobile water. However, the immobile water percentage was mostly higher in the AU group than in the CK group, whereas the reverse was true for the UA group. In the U3 and U2 groups (treated with ultrasound alone), the free water percentage dropped from 1.21% to 0.83%, and the immobile water rose from 94.76% to 95.50%. Therefore, longer high-intensity ultrasound treatment promotes water transfer to immobile water, enhancing the meat’s water-locking capacity. Sun et al. [34]. detected changes in water distribution in myofibrillar proteins by LF-NMR technique and found a shift from immobile to free water in the samples, proving that ultrasound action contributes to the changes in water distribution. They suggested that cavitation of ultrasound promotes unfolding of proteins and the exposure of hydrophobic groups, thus reducing the interaction between water and proteins. The opposite conclusion was reached in the present experiment, probably because of the difference in ultrasound intensity and time. Alternatively, Kang et al. [35]. proposed that the cavitation effect of ultrasound will expose the active region of the protein, enhance the ability of muscle fibers to bind water, and improve water retention properties. Therefore, the effect of ultrasound on the water retention properties of meat depends on the ultrasound intensity, time, and frequency.

The changes in the percentage of various water contents of abalone meat after heating are shown in Figure 2L. Among them, the U8 group had the highest immobile water proportion at 96.47%. At the same heating temperature, the immobile water proportion in the U group was higher than that in the B group, except for the B7 group, where the bound water proportion differed significantly from that in the U group. Meanwhile, the free water contents of both groups U7 and U8 were lower than that of group B, with significant differences. Therefore, there was a transfer from bound and free water to immobile water in group U. In the R group, the immobile water percentage was higher in R5 and R7 than in B5 and B7. However, the reverse was true for the other two groups. Moreover, the water transfer differed from that in the U group. Li et al. [36] used different time and temperature for steaming chicken breast meat to study its water retention capacity. They found that the amount of immobile water was higher in chicken breast meat steamed at 70 °C than at 60 °C in 60 min of steaming time, which was similar to the results of the present study. During heating, proteins gelatinize, allowing them to absorb more water, which flows back into the voids of the muscle fiber filaments [37]. The effects of ultrasound and enzyme treatments on the protein structure may accelerate this process, exhibiting a higher percentage of immobile water. The cavitation and mechanical effects of ultrasound on the myofibril structure result in the reorganization of myofibrillar proteins to form a complex and subtle protein network structure to hold more water [38].

MRI in Figure 3 and Figure 4 visualize the moisture content and distribution in the sample. Through the pseudo-color software processing, the color shades in the images represent the level of moisture content, with the deeper orange–red color indicating higher moisture content, and the deeper blue–green color indicating lower moisture content. Considering the differences in samples and parameter settings, there was no correlation between the color shades of images between raw and cooked meat. The lighter center and edge positions of the images in the experimental group of raw meat represented different degrees of moisture loss. The lighter color around the two enzyme-only groups (A3 and A5) indicated moisture loss from enzyme treatment alone. Ultrasound treatment resulted in a darker, more uniform reddish–orange color, particularly in AU53 and UA53. The darker areas of heated abalone meat were more uniform, suggesting that the heating treatment will promote the uniform distribution of water. The color gradually deepened with the increase in heating temperature, indicating the enhancement of the ability to fix water. The color of group U tended to be more orange–red compared with that of group B at the same temperature, which indicated the enhancement of the ability to fix water. However, the colors of R6 and R8 were yellowish–green, indicating that the water distribution was not uniform. Therefore, the difference in the order of ultrasound and enzyme treatment leads to the uniformity of water distribution and shows the difference in the percentage of fixed water and water status.

Figure 3. Effect of hydrogen proton density of abalone meat before and after ultrasound and enzyme treatment on treatment.

Figure 4. Effect of heat treatment on hydrogen proton density in abalone meat.

3.3. Cooking Loss

Cooking loss can indicate the structural integrity of the meat and reflect the water retention properties of the meat. As shown in Figure 1A, enzyme addition and increasing concentration caused the cooking loss of abalone meat to rise. Compared to the CK group, the cooking loss was increased by 0.56% and 1.43% in the A3 and A5 groups, respectively. Enzymes affect meat proteins by altering protein structure, degrading proteins, and destroying the original fibrous and reticular structures of meat, thereby increasing cooking loss and reducing water retention capacity [39]. Compared to the CK group, the U2 group was elevated by 0.87%, and the U3 group was reduced by 0.77%. Li K et al. [40] investigated the effect of different ultrasound time and salt addition on chicken mince and found that 40 min of ultrasound treatment increased the cooking loss of chicken mince by about 1% or so. Compared with the CK group, the AU32, AU33, AU52, and AU53 groups exhibited reductions in cooking loss of 0.10%, 0.37%, 0.23%, and 0.67%, respectively. In the UA group, the first three groups showed increases of 1.45%, 0.93%, and 0.17%, while the UA53 group had a 0.20% decrease. Figure 2A shows that the cooking loss of cooked meat gradually decreased as heating temperature increased. At the same heating temperature, the cooking loss in group U was 10.00%, 6.00%, 4.00%, and 4.33% lower than in group B. The cooking loss in group R was 10.00%, 7.33%, 4.67%, and 4.67% lower than in group B. Zou et al. [41] found that the ultrasonic power of 400–800 W could improve the water retention capacity of beef during the heating process. Improved water holding capacity after ultrasound may be due to an increase in the internal space of muscle fibers to hold more water [42].

3.4. Color

The total pigment content in meat and the chemical state of myoglobin affect meat color. Myoglobin can interact with other meat components, and changes in its concentration and redox state are among the most important factors influencing meat color [43]. As shown in Figure 1 and Figure 2G–I, enzyme treatment alone resulted in a decrease in L*. Ultrasound and combined treatments increased L*. The L* values indicate the lightness of meat, which correlates with its water content: higher immobile water content corresponds to a higher L* value. Combined with the increased proportion of immobile water observed in previous experiments, this explains the changes in L* values for both raw and cooked meat. The a* values represent the redness and greenness of the meat, which are mainly affected by the oxygenated myoglobin concentration. It was previously demonstrated that papain treatment further increased the amount of oxygenated myoglobin in meat by approximately 20% [44]. This could explain the rise in a* values, and it has also been shown that ultrasound treatment limits the production of oxygenated myoglobin [45]. The increase in a* values in this experiment may be attributed to the enzyme promoting a greater increase in oxygenated myoglobin content than the inhibitory effect of ultrasound, thereby increasing a* values. In cooked meat, the a* value gradually decreased with the rise in temperature, possibly because of protein loss due to temperature increase. Wang et al. [31]. also found a decrease in a* values when they used ultrasound-assisted frying of meatballs. The b* value represents the yellow and blue coloration of the meat, and oxidation and degradation of proteins affect the change in b* value. Oxidation of tryptophan and cysteine to produce a yellow substance also leads to an increase in b* values [46]. Ultrasound cavitation and enzymes cause cleavage of the spatial structure of proteins, while heat treatment also induces protein dissociation. This exposes tryptophan buried within proteins, which in turn triggers its oxidation—a process that may have contributed to the increase in b* values. Differences in a* and b* values between the AU and UA groups may be attributed to the reversed treatment order. In contrast, differences in L* values result from variations in water content and distribution.

3.5. pH

As shown in Figure 1 and Figure 2J, among abalone meat treated with enzyme and ultrasound, the pH reached a maximum of 6.58 in group AU53 and a minimum of 6.52 in group U3. Among them, the pH of group U2 was 6.54, which was higher than that of group U3, which was 5.52, and the prolongation of ultrasound time lowered the pH of the meat. Roobab et al. [47] applied ultrasound to chicken meat at 300 W, 500 W, and 800 W to study the effect on its physicochemical properties. They observed that pH decreases with increasing ultrasound power and duration. This is because the rapid collapse of microbubbles generated during the initial cavitation phase of ultrasound induces oxygen dissociation and hydrolysis, producing OH· and OOH· free radicals. These radicals release protons, thereby lowering pH. Their finding matches the results of this experiment. The group treated with both ultrasound and enzyme shows a gradual increase in pH. This is similar to the findings of Cao et al. [48]., who used a combination of papain and ultrasound to treat chicken breasts; the pH of the enzyme-only group was 5.75, which was significantly higher than that of the control group, which was 5.70. However, there was no significant change in pH after using a combination of ultrasound and enzyme in their study, which is similar to the results of the present experiments. This is because pH changes induced by the enzyme did not counteract free radical changes caused by high-intensity sonication. In cooked meat, pH was proportional to the increase in temperature. No significant differences were observed. This is in agreement with the study of Zhao et al. [49], who used ultrasound-assisted cooking of pork meatballs, which also showed no significant difference in pH.

3.6. Observation of Tissue Microstructure

Figure 5 and Figure 6 shows the scanning electron micrograph of abalone meat, which can visualize the microstructure of abalone meat muscle fibers. In this experiment, the myofiber network structure of the CK group had fine hollows. Enzyme treatment caused the network structure to collapse gradually. In groups A5 and A6, larger pores and fracture separation between fiber filaments were clearly observed. The ultrasound-treated groups U2 and U3 also consisted of pores, but the pores were finer and the fiber filaments were tighter than those in group A. In the groups AU32 to AU53, the muscle fiber filaments were tightly aligned, and the pores were even smaller. This indicates tightening of muscle fibers under the combined effect of ultrasound and enzyme, and indirectly on water fixation. In the UA group, the fibers in UA32 were loosely arranged. With increasing enzyme concentration and prolonged ultrasound time, the fibers tightened but remained more furrowed. Zheng et al. [50] applied 100 W low-intensity ultrasound to assist corned beef processing. They observed muscle fiber fragmentation, impaired structural integrity, and expanded interstitial spaces in the ultrasonic-treated beef. Similar phenomena were observed in the U2 and U3 groups of this experiment. Saengsuk et al. [51] treated ground pork using bromelain and found that the surface texture of the treated samples was smoother, while larger collagen fibers were still observed in the untreated samples. Both bromelain and papain are common enzymes used for meat texture improvement and have similar effects. In cooked meat, small holes were still observed in group B. In groups R and U, the fibers tightened and loose fibrils disappeared. In groups U7 and U8, the surface was smoother and flatter. However, in group R, cracks of varying sizes were still observed. The changes in muscle fiber filaments caused by the three treatments were important factors in the changes in water distribution and water holding properties.

Figure 5. Effect on microstructure of abalone meat before and after ultrasound and enzyme treatment.

Figure 6. Effect on microstructure of abalone meat after heat treatment.

Figure 7 and Figure 8 shows the optical micrographs of abalone muscle fiber tissues, and the fracture condition of abalone meat fiber filaments was observed. The fiber filaments in groups A3 and A5 were obviously fractured and arranged haphazardly. There were broken fiber filaments and some tightly arranged fibers in groups U2 and U3. In the AU group, the fiber filaments were tightly arranged. The AU32 group had inter-fiber pores. These pores reduced as ultrasonication time prolonged and enzyme concentration increased. The UA group was similar to the AU group but the inter-fiber pores were larger, especially in the UA32 and UA33 groups. The UA group was similar to the AU group, but the inter-fiber pores were larger, especially in the UA32 and UA33 groups. In the cooked meat group, as heating temperature increased, the fibers became densely and orderly arranged. This was most evident in the U8 group, whereas other groups still had pores of varying sizes.

Figure 7. Effect of ultrasound and enzyme treatment on abalone meat tissue fiber.

Figure 8. Effect of heat treatment on abalone meat tissue fiber.

3.7. TCA-Soluble Peptides

TCA-soluble peptides reflect the content of small molecular peptides, which in turn reflects the degree of hydrolysis of proteins under ultrasound and enzymes. Its higher content indicates higher degradation of proteins [52]. As shown in Figure 1 and Figure 2C, the addition of enzyme and its increased concentration led to a rise in TCA-soluble peptide content. Compared to the CK group, the A3 and A5 groups showed increases of 8.66% and 10.16%, respectively. In the ultrasound group, the U2 and U3 groups exhibited increases of 5.19% and 3.29% relative to the CK group. The contents of AU32, AU33, AU52, and AU53 groups increased by 6.22%, 5.99%, 5.78%, and 4.28% compared to the CK group. Gao et al. [53] used different protease inhibitors to inhibit histone L activity in minced silver carp. They found a significant decrease in TCA-soluble peptides, with a positive correlation between protease activity and TCA-soluble peptide content. In this experiment, direct application of protease to the meat inevitably led to an increase in TCA-soluble peptide content. In addition, the destruction of muscle fibers by ultrasound and enzymes caused the original protein structure to dissociate into small molecules. Consequently, the content of TCA-soluble peptides increased. In the cooked meat group, the TCA-soluble peptide content decreased with the prolongation of the heating temperature; B5 to B8 decreased by 11.39%, U5 to U8 decreased by 3.07%, and R5 to R8 decreased by 11.01%. Heat treatment gels the proteins, and during the gelation process, the interactions between the proteins lead to internal stabilization and reduced degradation, which affects the quality of the gel-type muscle foods [54]. This may also lead to a decrease in TCA-soluble peptide content.

3.8. BPB Binding Content

The degree of exposure of hydrophobic groups reflects changes in the hydrophobicity of the protein itself, allowing the assessment of protein surface activity and the determination of changes in the protein’s structure [55]. Changes in protein structure led to exposure of internal hydrophobic groups and increased BPB binding content [56]. As shown in Figure 1 and Figure 2F, all experimental groups were higher than the CK group, indicating that the effect of enzyme and ultrasound resulted in the alteration of protein structure. Groups A3 and A5, which were treated with enzymes only, showed a significant increase in BPB binding compared to the CK group, by 3.45% and 5.98%, respectively. The highest amount of BPB binding was found in group A5, which was 71.43 ± 0.17 μg. The protein was cleaved by the enzyme, and the internal hydrophobic groups were exposed. Using the combined action of enzyme and ultrasound, BPB binding content increased by 3.41%, 3.20%, 3.26%, and 3.42% in the AU group compared to the CK group, and by 3.51%, 3.42%, 3.68%, and 3.69% in the UA group, respectively. Sun et al. [34] investigated the binding effect between chicken myofibrillar proteins and spice compounds. They found that ultrasonication increased the amount of protein bound to BPB, with the maximum reached after 30 min of ultrasonication. However, there was no significant difference in the amount of BPB bound between the AU and UA groups. This may be due to protein aggregation [57] and the disruption of hydrophobic groups by enzymes. In the cooked meat group, the amount of BPB binding increased gradually with increasing heating temperature, from 35.90 ± 0.29 μg in B5 to 41.80 ± 0.44 μg in B8, from 40.66 ± 0.66 μg in U5 to 41.85 ± 0.17 μg in U8, and from 41.26 ± 0.04 μg to 42.02 ± 0.11 μg in R5. Wang et al. [32] cooked pork for different times and also found an increase in BPB binding from 44.23 μg to 113.27 μg. This is consistent with the results of the present experiment. This is likely because heating breaks hydrogen bonds and ionic bonds in proteins, which in turn causes protein chain breakage, conformational changes, and exposure of hydrophobic groups. In addition, the exposure of hydrophobic groups is associated with changes in the secondary structure of proteins, and the transition from α-Helix to β-sheet also affects the hydrophobicity of proteins.

3.9. Fluorescence Spectrum

Tryptophan and tyrosine groups are highly sensitive to the external environment. They are encapsulated within proteins. This helps maintain their quantum yields and fluorescence intensities. The fluorescence intensities of the emitted light can be analyzed by excitation of tryptophan and tyrosine groups in the samples, and the changes in the conformation of proteins can be analyzed [58]. As shown in Figure 9B, in the raw meat group, fluorescence intensity detection revealed that all experimental groups (A3-UA53) showed fluorescence quenching compared to the CK group. In the cooked meat group (B5-R8), the heat treatment destroyed the protein and thus led to a fluorescence burst, and the maximum fluorescence intensity decreased with the increase in heating temperature. It can be observed that longer ultrasound times and higher enzyme concentrations trigger more severe fluorescence quenching. Tryptophan groups are typically encapsulated in hydrophobic pockets within proteins. Enzyme-induced protein dissociation alters their spatial structure. The exposure of hydrophobic groups observed in protein hydrophobicity assays also indicates the disruption of these hydrophobic pockets. The two enzyme-treated groups (A3 and A5) exhibited the most severe fluorescence quenching. Here, proteins were cleaved by enzymatic action, leading to protein chain breakage and dissociation of the chromophore moiety. The disruption of hydrophobic pockets exposes tryptophan more to the external environment. Additionally, free radicals generated by sonication attack chromophore groups, which also contributes to fluorescence quenching [59]. The protein aggregation caused by high-intensity ultrasound will hide the tryptophan group and maintain a certain fluorescence yield. As a result, the fluorescence intensity of the sonicated U and AU groups was higher than that of the A group, and the fluorescence intensity was changed due to the involvement of enzymes.

Figure 9. Fluorescence spectra of myofibrillar proteins in cooked (A) and raw (B) abalone.

3.10. Total Sulfhydryl Groups and Free Amino Groups

Sulfhydryl and amino groups are side chain groups of proteins, and changes in their content reflect the extent of protein structural changes and the integrity of side chains. Total sulfhydryl groups refer to the sum of sulfhydryl groups exposed on the surface of protein molecules, and surface sulfhydryl groups of adjacent protein molecules can form disulfide bonds [60]. As shown in Figure 1 and Figure 2D, sulfhydryl content gradually increased with the addition and concentration of the enzyme, as well as with the application of ultrasound and the prolongation of treatment time. Compared to the CK group, the sulfhydryl content of the A3, A5, U2, and U3 groups increased by 5.00%, 6.06%, 1.83%, and 0.89%, respectively. Enzymatic and ultrasonic actions altered protein structure, and the unfolding of protein chains exposed more sulfhydryl groups. Thus, when ultrasound and enzyme were combined to treat abalone meat, the sulfhydryl group content increased further. Compared to the CK group, the sulfhydryl group content of the AU and UA groups increased by up to 4.14% and 4.02%, respectively. In cooked meat, the sulfhydryl content gradually decreased with the heating time, from 9.54 μmol/mg protein in B5 to 8.50 μmol/mg protein in B8, 8.82 μmol/mg protein in U5 to 8.08 μmol/mg protein in U8, and 8.59 μmol/mg protein in R5 to 7.83 μmol/mg protein in R8. The decrease in sulfhydryl content may be due to the oxidation of sulfhydryl groups in cysteine to disulfide bonds, which leads to the formation of intermolecular or intramolecular protein cross-links. In addition, proteins cross-link via disulfide bonds to form a dense network structure, which exacerbates muscle contraction and thereby affects water retention properties [61].

As shown in Figure 1 and Figure 2E, the free amino groups content of A3 and A5 increased by 5.26% and 6.06%, respectively, compared to the CK group. After ultrasound treatment, the free amino groups content of the U2 and U3 groups increased by 3.31% and 3.81% compared to the CK group. After the combined action of ultrasound and enzyme, the free amino groups content further increased. Compared to the CK group, the free amino groups content of the AU32, AU33, AU52, and AU53 groups increased by 5.56%, 5.06%, 5.11%, and 4.81%, respectively; that of the UA32, UA33, UA52, and UA53 groups increased by 5.37%, 4.66%, 4.61%, and 4.28%, respectively, while in the cooked meat group, there was no significant difference in the change of amino content. Analysis of sulfhydryl and free amino groups content shows that enzymatic and ultrasonic actions induce changes in protein structure and side chain groups. These changes affect the fiber integrity of abalone meat. Meanwhile, the oxidation of sulfhydryl groups to disulfide bonds strengthens protein cross-linking, which contributes to changes in hardness. Additionally, the tightened network structure enhances water retention capacity [62,63].

3.11. FT-IR

Structural changes in proteins are often determined using Fourier transform infrared spectroscopy. Figure 10 and Figure S1 in the Supplementary Materials show the FT-IR spectra of all sample sets with split-peak fitting of the 1600–1700 nm bands to calculate the percentage of protein secondary structure. In general, the amide I band of the FT-IR spectra 1600–1700 nm (indicating the C=O stretching/hydrogen bonding associated with -COO) contains variations in protein secondary structure and can be categorized into α-Helix (1650–1660 nm), β-Turn (1660–1670 nm), β-sheet (1600–1640), and Random coil (1640–1650 nm) [64]. In addition, FTIR spectra also include amide II band (1480–1575 nm), amide III band (1230–1300 nm), amide A band (3300 nm), and amide B band (2900–3000) [65]. In the cooked meat group, the amide band shifted to lower wave numbers (group R: 1651→1636 nm; group B: 1649→1639 nm; group U 1646→1638 nm). In the raw meat group, the amide I band showed the same trend as in the cooked meat group with increasing enzyme concentration and ultrasound duration, attributed to β-sheet formation. Bertram et al. [66] also observed a decrease in amide I wave number during heating. As shown in Figure 10A,B, in both the raw and cooked meat groups, the percentage of β-structures increased with increasing enzyme concentration, ultrasonication time, and heating temperature. Specifically, there was a shift from α-helix and random coil to β-turn and β-sheet. Ultrasonication alters the spatial entanglement of protein molecules, causing structural collapse, partial proteolysis, exposure of protein residues, and formation of hydrogen bonds. This leads to the shift from α-structures to β-structures [67]. β-sheet has a large surface area, which facilitates cross-linking between protein molecules [68]. This could explain the enhanced water retention properties. In addition, protein loss can result from the attack of proteins by free radicals generated by high-intensity ultrasound, enzymatic degradation of proteins, and heat treatment. Wang et al. [69] reported that the decrease in α-structure and increase in β-structure may be due to oxidative loss of proteins, and α-helix content and surface hydrophobicity showed a negative correlation, which is consistent with the structure of the present study.

Figure 10. Secondary structure occupancy (A,B) of proteins in raw abalone (A) and heat-treated abalone (B).

3.12. SDS-PAGE

As shown in Figure 11, proteins were separated using SDS-PAGE, and changes in protein molecular weight were observed. After enzyme treatment, the MHC bands around 270 kDa disappeared in the A3 and A5 groups, indicating that proteases degraded MHC. Meanwhile, the intensification of bands between 50 and 90 kDa was observed, confirming the formation of small fragments. In the A5 group, the degradation of large molecular weight bands was greater than in the A3 group. Notably, the loss of bands in the 40–53 kDa range was evident, indicating that higher enzyme concentrations caused more severe protein degradation. The changes in the 270 kDa bands in the U2 and U3 groups after ultrasound treatment did not change compared with those in the CK group. However, the MHC bands in the AU group were slightly lighter compared with those in the CK group, especially the most obvious in the AU53 group. In addition, the shift of bands toward higher molecular weights around 20 kDa and the deepening of bands at 90 kDa further indicated that proteins were transferred to higher molecular weight fractions. When analyzed in conjunction with the changes in the bands in the enzyme-treated group, high-intensity sonication may have led to the polymerization of proteins. The disappearance of the 270 kDa band and the deepening of low-molecular-weight bands in the UA group may be attributed to protein degradation induced by enzyme treatment. In the cooked meat group, the protein molecular weights in the U5–U8 groups were higher than those in the B5–B8 groups (which underwent direct heating). The 270 kDa bands were distinct in the U5–U8 groups, whereas all proteins with 270 kDa bands in the B groups were degraded. However, the protein molecular weights in the B groups gradually increased with rising heating temperatures, indicating that temperature influences protein aggregation. This trend was similarly observed in the R group. In addition, in the range of 40~53 kDa, the protein bands of group B5~B7 were severely lost. Consequently, the molecular weight increases, accompanied by a decrease in sulfhydryl content. Previous studies have demonstrated that heat treatment induces protein gelation, promotes disulfide bond formation, and strengthens protein aggregation [70]. The results of Qi et al. [71] in the study of stewed chicken meat were also similar to the present study. Taken together, high-intensity ultrasound induces protein polymerization and compensates for protein loss caused by enzyme treatment. This is one of the factors contributing to changes in the water retention properties of the meat. In contrast, ultrasound followed by enzyme treatment results in more severe protein degradation. It impairs the formation of protein networks and slows down the gelatinization process during heating.

Figure 11. SDS-PAGE of abalone meat proteins before and after ultrasound and enzyme treatment and after heat treatment; Marker from top to bottom: 270, 165, 130, 90, 68, 53, 40, 30, 20, 13, 8 kDa.

4. Mechanism of Textural Property Changes in Abalone Meat

Hardness was correlated with the water-holding capacity of meat and the integrity of fibrous tissue. Ultrasonic and enzymatic treatments increased the percentage of immobile water in abalone meat. Enzymes cleaved proteins and muscle tissue fibers, thereby enhancing the plasticity of abalone meat. The mechanical effects of high-intensity ultrasound induced movement in abalone muscle tissue fibers and promoted protein–protein interactions, which tightened fiber filaments and thereby contributed to water retention. Abalone meat is inherently tough and requires effort to chew. When treated with protease first, muscle fibers are relaxed, proteins are cleaved, and internal hydrophilic groups are exposed to attract water molecules. Subsequent ultrasonic treatment tightens the fiber filaments, fixing the attached water and thus increasing the proportion of immobile water. Conversely, high-intensity ultrasound treatment applied first also contributes to water retention to a certain extent. However, the contraction of already tight fibrous tissue under ultrasound may increase hardness, while the cavitation effect of ultrasound can break fiber filaments, resulting in reduced hardness. This study demonstrated that high-intensity ultrasound induced protein aggregation in abalone meat. Microscopic observations also revealed the coexistence of contracted fibrous structures and broken fiber filaments, which collectively influenced the hardness of abalone meat. Subsequent enzyme treatment caused the breakage of contracted fiber filaments, resulting in the loss of fixed water. Meanwhile, hydrophilic groups exposed by protein cleavage attracted water molecules but failed to retain them effectively. While this reduced hardness, it suggests that fiber damage caused by proteases predominated. In heat treatment, the hardness of abalone meat decreased with increasing temperature, and proteins gelatinized due to heat treatment, forming a dense structure, which facilitated the uniformity of the meat network and thus water retention [72]. However, the excessive aggregation of proteins forms large insoluble material, which affects the continuous protein gel network, resulting in reduced structural properties [73]. In previous studies, kinetic modeling has been used to study the effect of heat treatment on meat quality [74]. Therefore, in order to obtain more accurate results, similar methods can be used to study the effects of temperature and time changes on the textural properties of abalone meat.

5. Conclusions

Both raw abalone meat subjected to ultrasound and enzyme treatments and cooked abalone meat subjected to heat treatment exhibited reduced hardness. The sequence of ul-trasound and enzyme application affected meat hardness and water-holding capacity, with meat hardness showing an inverse correlation with the percentage of immobile water. Protein cleavage in treated raw meat and gelation in heated cooked meat influenced the content of TCA-soluble peptides. Color changes were associated with protein degradation, loss of side chain groups, and altered water-holding capacity. Prolonging ultrasonic treatment time, increasing enzyme concentration, or elevating heating temperature reduced the hardness of abalone meat and altered protein structure. Both treatment sequences exhibit the conversion of α-helices to β-turns and β-sheets. As the heat treatment temperature increases, protein molecular weights increase, and tissue fibers become tightly arranged.