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. 2023 Feb 4;94:106318. doi: 10.1016/j.ultsonch.2023.106318

Underlying formation mechanisms of ultrasound-assisted brined porcine meat: The role of physicochemical modification, myofiber fragmentation and histological organization

Guofeng Jin 1,1, Yuanyi Liu 1,1, Yan Zhang 1, Chengliang Li 1, Lichao He 1, Yuemei Zhang 1, Ying Wang 1, Jinxuan Cao 1,⁎
PMCID: PMC9932671  PMID: 36753829

Highlights

  • •

    UAB induced stronger muscle protein denaturation and proteolysis of pork than SB.

  • •

    UAB could enhance myofiber fragmentation and disintegration of muscle tissues.

  • •

    Muscle showed higher fibril and perimysium swelling capacity after UAB treatment.

Keywords: Ultrasound; Brining; Meat; Postmortem proteolysis; Fiber fragmentation, Microstructure

Abstract

Ultrasound treatment has been a good hurdle technique for meat curing processing, where both physical and chemical consequences can be involved towards final quality of obtained products. However, the specific correlation between ultrasound parameters and muscle fiber fragmentation and myofibrillar microstructural changes during curing deserve further evaluation. In present study, we comparatively studied the effect of ultrasound-assisted brining (UAB) and static brining (SB) on the muscle proteolysis events and microstructural/morphological variation of porcine meat as well as the physicochemical indices and histological characteristics. The results showed that UAB (20 kHz, 315 W for 1 h) could markedly enhance the muscle proteolysis with higher free-/peptide-bound alpha-amino-nitrogen (α-NH2-N) content (P < 0.05) than SB treatment and greatly improved the fragmentation of muscle fiber tissues of cured meat. Meanwhile, UAB processing favored more opening structures of myofibrillar proteins with more hydrophobic groups being exposed. The quantitative histological analysis revealed that, compared with SB treatment, UAB could significantly increase the gap between muscle fibers and the swelling of the perimysium (P < 0.01), proving an efficient curing process with better textural and water holding properties.

1. Introduction

Brining is an important meat processing technology. During brining, the meat is immersed in a salt solution where it absorbs extra liquid and salt. Various combinations of salting procedures (e.g., injection and/or brining or pickling, followed by dry salting) have been practically used to ensure different degree of protein aggregation and moisture loss at initial stage of curing. Meanwhile, the penetration of NaCl into meat could result in many physicochemical changes of the meat matrix [1]. Common to all curing methods is that NaCl diffuses through the meat matrix and the diffusion rate depends on the local concentration gradient. The diffusion of NaCl into the meat under normal conditions (traditional static brining) is usually time-consuming due to the complex matrix of meat. So, in many cases, the brining process is often complemented by some other treatments, such as tumbling, high pressure, and ultrasound, which could cause cellular disruption of meat tissue [2].

Ultrasound principally acts by generating bubble cavitation in the biological matrix, and the consequent cavitation effects are the major cause of physicochemical reaction of muscle during processing. Indeed, an improved meat tenderness with desirable final quality can be induced by ultrasonic treatment by breaking the weak interactions between molecules, resulting in the disruption of muscle fiber fracture [3], [4]. As a new green and environmentally friendly technology in recent years, ultrasonic-assisted brining has been deeply studied and widely used in the brining process and texture improvement of meat products [5]. Compared with other curing techniques, ultrasound treatment can significantly improve the structure of cured meat and promote the transfer of sodium chloride in the muscle, but also induce many chemical reactions that affect the color and tenderness of the meat, which is of great significance to the curing industry [6]. Guo et al. [7] proved that the simultaneous ultrasound treatment with the different frequencies hastened NaCl diffusion into porcine tissue and improved the color and tenderness of the cured pork, meanwhile favored the pickling absorptivity and water-holding capacity. Furthermore, in our previous study, the results showed that the UAB (20 kHz, 350 W for 1 h) could significantly increase the oxidative susceptibility of muscle proteins, which consequently improved the water holding capacity (WHC), color and textural properties (hardness, springiness and chewiness) of the cured meat samples compared to the static brining (SB) samples (P < 0.05) [8]. Alves et al. [9] also reported that the use of ultrasound (25 kHz, 128 W) treatment in manufacture of dry fermented sausages could significantly affect the proteolysis and formation of the volatile flavor compounds derived from lipid oxidation during the storage. Bao et al. [10] showed the ultrasound pretreatment (300 W, 20 kHz) could significantly decrease the hardness of dry-cured yak meat by enhancing the muscle proteolysis. For these phenomenon, previously studies have underlined that ultrasound can positively interfere with the activity of protease and lipase, which can affect the muscle proteolysis and lipolysis [11], [12]. And the ultrasonic activation of brine could increase the hydration level of muscle proteins, which also affects the hydrolysis of the proteins [13]. Kang et al. [14] revealed that the power ultrasound-assisted curing with 20 kHz frequency processing could improve WHC and tenderness of the beef, meanwhile moderate oxidation induced myosin polymerization which positively contributed to the water retention of muscle.

Moderate proteolysis and myofiber breakage are conductive to obtain desirable quality of cured meat. As far as we know, in the past decades, many studies have indicated that both the individual treatment of brining and ultrasound could obviously affect the muscle proteolysis and myofiber structure changes, resulting in the changes of textural and flavor quality [6], [15]. Du et al. [3] showed the ultrasound treatment-assisted tenderization could significantly improve the tenderness of chicken gizzard by weakening the mechanical strength of connective tissue and also improve the degree of myofibril fragmentation. However, few studies have assessed the combined effect of ultrasound-assisted brining on cured meat quality concerning the mechanistic changes in muscle tissues and the profiles of muscle proteins hydrolysis to form amino acids and peptides.

Thus, the objective of this study was mainly to investigate the effect of UAB on the muscle protein hydrolysis and microstructure changes during curing process. The muscle protein hydrolysis degree (proteolytic index, PI), characteristic proteolysis intensity (α-amino nitrogen, peptide nitrogen), myofibrillar protein structure (surface hydrophobicity, intrinsic fluorescence), muscle fiber breakage (myofibril fragmentation index, MFI), and the muscle histological properties as well as microstructure of the cured meat were explored to better explain the mechanism of ultrasound-assisted brining-induced changes of WHC and tenderness.

2. Materials and methods

2.1. Sample preparation and brining treatment

Fresh biceps femoris (BF) from the same pig carcass (Taihu × Duroc × York crossbred male pig 9-month-old, weighing 100 ± 2 kg) were purchased at a local slaughterhouse (Wuhan, China). The pieces selected had a pH of 5.8 ± 0.30, which was measured in-situ by a pH-meter (FiveEasy28, Mettler-Toledo Instruments Co., Ltd, Shanghai, China) at three different points along the muscle avoiding fatty areas. the visible fat and connective tissues were trimmed off from the muscles. A total of 9 parallelepiped shaped samples (length 60 × width 40 × thickness 40 mm) with an average weight of 350 ± 2 g were obtained from the central part of biceps femoris pieces using a sterilized knife along the fiber direction. Before brining, samples were vacuum packaged in polyamide/polyethylene (PA/PE) bags and kept frozen at − 20 °C.

Then, these packaged strips were slowly thawed at 4 °C for 24 h and randomly divided into three groups (three strips per). Apart from the control samples those were kept at 4 °C for 3 d, the other two groups were treated by static brining (SB) and ultrasound-assisted brining (UAB), respectively. For SB, the samples were immersed in 6 % NaCl solution (1:5 w/v) at 4 °C for 3 d without any agitation, while UAB samples were marinated in 6 % NaCl solution (1:5 w/v) with 1 h ultrasonic treatment (20 kHz, 350 W, 31.02 W/cm2) (SCIENTZ-IID, Ningbo Xinzhi ultrasonic technology Co., ltd, Zhejiang, China) at the beginning of the whole brining process, and other conditions were consistent with SB samples. The actual ultrasonic intensity was determined according to the method of Kang et al. [15].

2.2. Proteolysis index

The determination of total nitrogen (TN) used the Kjeldahl method based on the standard method of International Standard Organization (ISO 937:1978) [39]. Digested 2.0 g of each muscle sample using a hot digester and then cooled to the room temperature, tritrated using automatic kjeldahl apparatus (Kjeltec™ 2300, Foss, Denmark). The measurements were triplicated. TN values were expressed in the unit of mg N/100 g.

Non-protein nitrogen (NPN) was determined according to the method of Harkouss et al. [16] with slight modification. Five grams of sample was minced and treated with 25 mL of 10 % cold trichloroacetic acid (TCA). The mixture was homogenized (Ultra Turrax T18 basic, IKA Werke GmbH & Co. KG, Baden-Württenberg, Germany) for 60 s (18,000 rpm) in ice bath, and thereafter stood for 12 h at 4 °C. The homogenate was centrifugated (5,000 g × 5 min, 4 °C) and filtrated, the supernatants were collected, and transferred to a 50 mL volumetric flask. 10 mL of the prepared samples were determined following the TN determination method. The measurements were performed in triplicated. NTN was expressed in the unit of mg N/100 g.

The proteolysis index (PI) was defined as the percentage of NPN in TN of the sample.

2.3. Free and peptide bound alpha-amino-nitrogen (α-NH2-N)

The free and peptide bound α-NH2-N were determined following the method of Oddy [17] with slight modifications. One gram of meat samples was homogenized in 20 mL of 0.6 mol/L perchloric acid (PCA). The homogenate was centrifuged and filtered. For total α-NH2-N analysis, 2 mL of PCA extract was mixed with 5 mL of HCl (8.4 M) and incubated at 100 °C for 24 h for hydrolysis. After that, the hydrolysate solutions were neutralized and diluted to 50 mL. One millilitre of a buffer solution, containing 0.5 % of ninhydrin (pH 5.8) and 100 μL of 50 % ascorbic acid solution, was added to either 0.2 mL of the PCA extract with 0.8 mL of distilled water (for free α-NH2-N) or 1 mL of the neutralized hydrolyzed extract (for total α-NH2-N). A solution containing 1 mL of distilled water, 1 mL of 0.5 % ninhydrin buffer solution (pH 5.8) and 100 μL of 50 % ascorbic acid solution was used as a blank. All mixtures were incubated for 20 min in boiling water, cooled to room temperature, and then added 5 mL of a 60 % ethanol solution, measured the absorbance at 570 nm. Results were calculated as mg α-NH2-N/100 g of dry matter according to reference method ISO 1442–1973, by comparing with solutions of ɩ-leucine of known concentration. A standard was used and treated the same way. The peptide-bound α-NH2-N contents were calculated by subtracting the free α-NH2-N from the total α-NH2-N.

2.4. Myofibrillar fragmentation index (MFI)

The MFI was determined according to the method of Poljanec et al. [18] with some modifications. Two grams of chopped sample of muscle tissue was homogenized twice at 10,000 g for 30 s each in intervals of 10 s in 40 mL of 4 °C extraction buffer (20 mM phosphate buffer, pH 7.0 containing 100 mM KCl, 11.2 mM K2HPO4, 8.8 mM KH2PO4, 1 mM EGTA, 1 mM MgCl2 and 1 mM sodium azide). The homogenate was centrifuged at 12,000g for 15 min at 4 °C, and the supernatant was discarded. The pellet was resuspended in 20 mL of extraction buffer and recentrifuged at 4,000 g for 15 min at 4 °C. The precipitate was resuspended in 10 mL of MFI extraction buffer and filtered through a polyethylene strainer to remove connective tissue and debris. The concentration of the filtrate, which was regarded as myofibrillar protein (MP) solution, was diluted to 0.5 mg/mL. The absorbance at 540 nm of the myofibrillar suspensions (0.5 mg/mL) was read in duplicate using an ultraviolet/visible spectrometer. The MFI was obtained by multiplying the mean value of absorbance reading by 200.

2.5. Intrinsic fluorescence spectra

The intrinsic fluorescence emission spectra of myosin were determined using a fluorescence spectrophotometer (RF-5301PC, Shimadzu, Japan). A certain amount of the extracted myosin protein samples were diluted with 5 mM phosphate buffer (0.6 M NaCl, pH 7.0) to a final concentration of 1 mg/mL. The emission spectra of tryptophan residues in protein solutions were then recorded ranging from 300 to 500 nm with the excitation wavelength established at 290 nm. The relative intrinsic fluorescence at emission maximum for assayed samples were obtained by subtracting the background signal from treated group.

2.6. Surface hydrophobicity

The method of Alizadeh-Pasdar et al. [19] was used to determine the surface hydrophobicity of myofibrillar protein with slight modification. Appropriate amount of myofibrillar protein was diluted to 0.5–5 g/L in phosphate buffer solution (pH 7.0 containing 0.01 M K2HPO4) according to 2-fold gradient series. For each protein concentration, 2 mL of each protein concentration was taken and reacted with 20 μL of 8 mM ANS (pH 7.0 containing 0.01 M K2HPO4) for 10 min. The fluorescence intensity of each protein concentration was measured under the condition of excitation wavelength 370 nm and emission wavelength 490 nm, and the reaction tube without ANS was used as blank. The surface hydrophobicity of protein was plotted with different protein concentration and corresponding fluorescence intensity, and the slope of regression fitting curve was used to represent the hydrophobicity of protein surface.

2.7. Muscle histology observation by hematoxylin and eosin (HE) staining

The muscle samples of different treatment groups were cut into pieces about 1 cm × 1 cm × 0.4 cm. The piece samples were fixed with 4 % paraformaldehyde at 20 °C for 2 h and embedded in paraffin blocks. After making frozen section, the tissues were stained with HE. The microstructure was observed under a fluorescence inversion microscope (DMIL-20512401, Germany) at 10 ×, 20 × and 40 × magnification, respectively.

2.8. Scanning electron microscopy (SEM)

The landscape and micro-morphology of the assayed tissues were imaged by means of a scanning electron microscope (S-4800 FE-SEM, Hitachi, Japan) as described by Pan et al. [8]. The muscle sample was cut into cubes with the side length of 2 mm and lyophilized in a freeze-dryer to complete dryness. The lyophilized samples were then gold-coated using an ion sputtering coating instrument, and an electron beam was generated in the SEM accelerated to 1–30 kV voltage. Samples were subjected to instrumental observation at 500 × and 1,000 × magnification, respectively.

2.9. Cross sectional area of muscle fibers and perimysium thickness

The cross-sectional area of muscle fibers and perimysium thickness of different muscle samples were determined by five SEM photographs obtained at 100 × magnification using individual muscle tissue. For each muscle tissue, five different microscopical fields were sampled. The Multi Scan Base v.13 computer image analysis software was used to measure the fiber cross-sectional area and the perimysium thickness. Five measurements of cross-sectional area and perimysium thickness were made on each image. The mean value of the 25 measurements was included in the statistical analysis.

2.10. Statistical analysis

All the experiments were in triplicate (n = 3) from three individual strips for each group, three determinations were included for each assay. The obtained results were statistically analyzed by SPSS statistical software (version 19.0), and shown as mean and standard deviation (SD). One-way analysis of variance (ANOVA) (P < 0.05, post-test Ducan) was performed to determine significant difference of datasets within the effect of brining methods (control group, SB and UAB samples). A mixed model was used to estimate the response of the measured parameters to the experimental factors such as ultrasonic treatment (as fixed term) and salt content (as random term regarding the source of meat pieces). Origin 2018 software (OriginLab, Northampton, Massachusetts, USA) was used for figure plotting.

3. Results and discussion

3.1. Effect of ultrasound-assisted brining on proteolysis index of cured meat

The changes of proteolysis index of porcine meat by ultrasound-assisted brining and static brining were shown in Fig. 1. Compared with fresh pork, the proteolytic index of SB and UAB sample increased significantly (P < 0.05), particularly for ultrasound-assisted brining group (P < 0.01). Zhang et al. [20] characterized the modifications of the proteome for unsmoked bacon following ultrasound treatment and identified a total of 137 differentially abundant proteins of bacon samples, most of which (81.02 %) showed a significant upregulation after ultrasound. These representative results could be due to the mechanistic action caused by cavitation upon ultrasonic treatment which probably leads to the rupture of tissue cells and the release of endogenous proteases [8], [21]. Endopeptidases (e.g., cathepsins and calpains) can degrade myofibril structures by cleaving the peptide bonds inside the proteins, leading to the generation of macromolecular polypeptides [20]. Wang et al. [21] found that ultrasound treatment induced increased proteolysis bovine semitendinosus (ST) muscle during the postmortem storage and improved the tenderness by modulating the activation of calpain and protein degradation. Consequently, muscle proteins can be continuously hydrolyzed into peptides, free amino acids and other flavor substances and their derivates, forming the flavor system of cured meat [22], [23].

Fig. 1.

Fig. 1

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Effects of ultrasonic-assisted brining (UAB) and static brining (SB) on PI.

3.2. Evolution of α-NH2-N and peptide nitrogen as affected by ultrasound-assisted brining

On the other hand, the free and peptide-bound α-NH2-N are regarded as efficient examiners to reflect the susceptibility of proteolysis in meat system [24]. Their contents in static brining group and ultrasound-assisted brining group were significantly higher than those in fresh pork (P < 0.05) (Fig. 2), which was in line with the result of proteolytic index (Fig. 1). The formation of α-NH2-N is usually accompanied with the collapse of primary structure of the muscle proteins and the subsequent cleavage of peptide bonds and the increase of free amino groups [21]. Following accessible hydration of skeletal muscle proteins during brining, the additional ultrasonic treatment and cavitation effect might further promote protein degradation where the local high temperature and high pressure could be involved [5], [23]. Noteworthy, higher contents of free and peptide-bound α-NH2-N are able to improve the organoleptic properties and oxidative stability of brined meat, which could be affected by some representative factors such as protease inhibitors and microenvironmental sequence/residue diversity [24], [25].

Fig. 2.

Fig. 2

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Effects of ultrasonic-assisted brining (UAB) and static brining (SB) on free α-NH2-N (a) and peptide nitrogen content (b).

3.3. Changes in myofibril fragmentation index (MFI) of brined meat

The myofibril fragmentation index (MFI) represents the integrity of myofibril and reflects the tenderness of meat [26]. The higher MFI value usually indicates damaged structures of myofibers and fibril subunits [21]. As depicted in Fig. 3, both static brining and ultrasound-assisted brining significantly increased the myofibril fragmentation index of porcine meat compared with control sample (P < 0.05), but the degree of enhancement toward fragmentation upon ultrasound treatment was the highest. Hence, the mechanistic effects of ultrasound-induced cavitation could produce intense physical forces such as shear forces, shock waves and turbulence product in liquid mediums, which may have dissociated muscle fibers with extensively high hydrolysis events experimentally observed [5], [21].

Fig. 3.

Fig. 3

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Effects of ultrasonic-assisted brining (UAB) and static brining (SB) on MFI.

3.4. The properties of endogenous fluorescence of myofibril of brined meat

The fluorescence pattern of endogenous tryptophan is often used to characterize the dynamically complementary features of hydrated proteins, which depends on the hydrophobic environment with protein unfolding and refolding process [27]. Fig. 4 revealed that the endogenous fluorescence intensity at an emission wavelength of ∼ 340 nm was significantly enhanced by brining treatment (SB and UAB) compared to the fresh pork (P < 0.05), particularly following UAB treatment. Hence, more hydrated structures of endogenous tryptophan have been extensively formed though their oxidative loss and quenching also take an integral part in the entire refolding course following intra-/inter-molecular interaction [24], [28].

Fig. 4.

Fig. 4

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Effects of ultrasonic-assisted brining (UAB) and static brining (SB) on protein intrinsic fluorescence.

3.5. Effect of ultrasound-assisted brining on surface hydrophobicity of brined meat

Surface hydrophobicity, as an important indicator to reflect the protein physical/chemical stability, usually depends on the diversity in exposed residues, flexible structure of globular proteins, and deamidation status following biochemical courses [29]. In general, the partial expansion of the animal proteins leads to the exposure of hydrophobic amino acid residues from buried structures, resulting in increased surface hydrophobicity [30]. It can be seen from Fig. 5 that the protein surface hydrophobicity of static brining group and ultrasound-assisted brining group increased significantly compared with fresh pork (P < 0.05), and the overall trend of ultrasound-assisted brining group showed a significant increase (P < 0.05). Accordingly, the structural instability of cured meat protein should be much favored by ultrasonic treatment than the static brining. During the curing process driven by salt diffusion, reasonably, the muscle tissue reached an enhanced proteolysis and broadens the protein-dense network structure, which could result in the exposure of hydrophobic groups [8], [31].

Fig. 5.

Fig. 5

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Effects of ultrasonic-assisted brining (UAB) and static brining (SB) on protein surface hydrophobicity.

3.6. Effect of ultrasound-assisted brining on histology of brined meat

Toward clearer understanding of the integrity of muscle tissue following brining, the muscle sections stained by HE were examined. As shown in Fig. 6, it was obvious that the fresh pork had cylindrical muscle fibers with the same cross-sectional area, and the myofibril bundles were completely arranged and parallelly organized with each other. However, ultrasound-assisted brining treatment and static brining treatment led to a certain degree of deformation of myofibril bundles, the relative increase of gaps between fibers as well as the fiber cross-sectional area. After ultrasound-assisted treatment, interestingly, the perimysium structure was further destructed and more obvious microfissures were identified between interspaces of myofibrils. Meng et al. [32] observed that the myofibril interval of bovine meat was increased and distributed unevenly during stewing. The fiber swelling following diffusion of salt ingredients are specific to ion-binding course which is concentration dependent [33]. An appropriate increase of electrostatic repulsion between muscle fibers and fiber gap (binding of chloride ions (Cl−) to protein filaments) during hydration process should be conducive to water retention of final brined product [34]. The water molecules are mainly bound by dipole-ion interaction between muscle fiber proteins. Therefore, an undesirable electrostatic repulsion can occur due to the substantive reduced net charge of protein molecules between fibrillar structures which usually leads to poor water retention [35]. Pan et al. [8] reported that, ultrasound-assisted brining (20 kHz, 350 W for 1 h) effectively disrupted the fibrous structures of porcine biceps femoris and gave rise to more noticeable swelling of muscle microstructures following the ultrasonic cavitation, meanwhile accelerated mass transfer and recovery of water mobility/population nearby myofibrillar network.

Fig. 6.

Fig. 6

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Histological profile (HE stain, 10×, 20× and 40× magnification) of brined meat upon ultrasonic-assisted brining (UAB) and static brining (SB) treatment.

3.7. Effect of ultrasound-assisted brining on myofibril cross-sectional area and perimuscular membrane thickness of cured meat

Muscle tissue is composed of a large number of myofibril bundles and the layer of perimuscular membrane is wrapped around the myofibril to maintain the structural stability. The thickness and water content of the myofibril affect the tenderness and texture of the muscle tissue [36]. Fig. 7 showed the visible changes of myofibril cross section and perimuscular membrane shape, area and thickness in each treatment group. It can be seen from Fig. 7A that the myofibril cross section of UAB samples was more regular and the surface of perineurial membrane appeared to be smoother than that of SB samples. Fig. 8 further proved that, unlike the static brining group, the myofibril of the ultrasonic treatment group was neatly arranged, and more well-fine connective tissue between the fibers was retained. We speculated that salt penetration was slower during static brining, and a higher scale of sodium chloride was distributed on the tissue surface, which led to a major difference in salt concentration around the tissue. Particularly for the perimuscular membrane which is immobilized water-abundant [37], the shifts in the protein isoelectric point and protein denaturation may occur due to close proximity to charged surfaces mainly under the binding action of Cl− during the curing process. As a result, the larger gap of salt ions intensity between the surface and internal section of the perimuscular membrane should be putatively responsible for higher degree of deformation in perimuscular membrane, thus showing an irregular morphology being wrinkling and curling on the surface of myofibril.

Fig. 7.

Fig. 7

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Evolutional cross-sectional pattern of fiber tissue by microscopy (a) and the quantitative cross-sectional area of myofibrils (nm2) and thickness of membrane perimuscularis (nm) (b) upon ultrasonic-assisted brining (UAB) and static brining (SB) treatment. Microscopic observation was done at 500× and 1000× magnification, respectively.

Fig. 8.

Fig. 8

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Effects of ultrasonic-assisted brining (UAB) and static brining (SB) treatment on the morphology of myofibrils and perimuscular membranes. Microscopic observation was done at 500× and 1000× magnification, respectively.

As quantified results, Fig. 7B-C showed that the myofibrillar cross-sectional area and thickness of perimuscularis of static brining group and ultrasound-assisted brining group increased significantly compared with untreated sample (P < 0.01). These phenomena could be involved with different causes: i) the neutralization of positively charged amino acid groups on the surface of myofibrillar proteins can be tailored by either Cl− or hydrolysis/rupture of proteins under the action of enzymes (e.g., activation of cellular protease system) and ultrasound during the curing process [1], [8], [14]; ii) the mechanistic action of ultrasound shows good potency in stabilization of immobilized water located around perimuscular membrane which makes the fiber swollen in relatively compact manner. Similar evidence of such effect have been observed in the cases of ultrasound-assisted enzymatic tenderization [37] and other mild cooking process of meat (e.g., steaming) [38]. Therefore, a desirable expansion of swelled myofibrils, water state and physical barrier for salt diffusion could be in favor of the texture, tenderness and other edible properties of these muscle foods.

4. Conclusion

We confirmed in present study that moderate ultrasound-assisted brining (20 kHz, 350 W for 1 h) could significantly promote the myofiber fragmentation and disintegration of muscle tissues by effectively accelerating the transfer of sodium chloride toward hydrated protein interface. Microscopic results revealed that ultrasound treatment promoted the fracture of myogenic fibers which contributed to higher myofibril fragmentation index, and the cross-sectional area as well as fiber interspace, implying a reasonably good textural properties of brined porcine meat. Accordingly, compared to static brining, a more extensive protein degradation of brined meat occurred following ultrasound treatment owing to higher fibril swelling capacity and reducible fluctuation of section between perimuscular membrane where both cavitation effect and the release of cellular protease probably made a difference. However, more detailed nutritional value of final products particularly the digestive profile at molecular level upon in vitro and in vivo condition require further evaluation.

CRediT authorship contribution statement

Guofeng Jin: Project administration, Funding acquisition, Supervision, Methodology, Conceptualization, Writing – original draft, Writing – review & editing. Yuanyi Liu: Investigation, Software, Validation, Writing – original draft, Writing – review & editing. Yan Zhang: Investigation, Software, Validation, Writing – original draft. Chengliang Li: Software, Validation, Formal analysis, Writing – review & editing. Lichao He: Software, Validation. Yuemei Zhang: Validation, Formal analysis. Ying Wang: Software, Validation. Jinxuan Cao: Project administration, Supervision, Funding acquisition, Writing – review & editing.

Declaration of Competing 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.

Acknowledgement

This research was supported by the National Natural Science Foundation of China, China (No. 31871824) and Fund of the talent introduction start-up project, Beijing Technology and Business University, China (No. 19008022334). The authors also appreciated the financial support provide by the fund of Cultivation Project of Double First-Class Disciplines of Food Science and Engineering, Beijing Technology & Business University (No. BTBUYXTD202202).

Data availability

Data will be made available on request.

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