The effect of ultrasound treatments on the tenderizing pathway of goose meat during conditioning

The effect of ultrasound treatments on the tenderizing pathway of goose meat during conditioning ABSTRACT In order to figure out the effect of ultrasound treatments (UT) on the tenderizing pathway of goose meat, breast muscles of 32 Eastern Zhejiang White Geese were treated with different ultrasound powers (the control, 300 and 600 W) at 40 kHz for 30 min. Shear force, cooking loss, myofibrillar fraction index (MFI), particle size, morphological analysis of actin filaments and the levels of F-actin and G-actin were investigated during 168 h storage. Results showed that 600 W group had the lowest shear force and cooking loss at 24, 48, 96 and 168 h among treatments, while 300 W UT decreased shear force and cooking loss compared to the control. UT increased MFI and induced myofibrillar small particles (D3,2) by disrupting actin filaments for myofibril and transforming of F-actin to G-actin compared to the control. We concluded that UT tenderized goose meat by fragmenting actin filaments and myofibrillar fraction. INTRODUCTION Tenderness is one of the most important quality parameters affecting consumer satisfaction and the positive perception of meat products (Miller et al., 2001). It can be improved by various physical, chemical and biochemical methods, such as electric stimulation, high-pressure shockwaves, tumbling, calcium marination and plant proteases (Jung et al., 2000; Hwang et al., 2003; Alvarado and Sams, 2004; Naveena et al., 2004; Feng et al., 2017; Li et al., 2017). It was demonstrated that the mechanism of tenderization is related to the destruction of myofibrils (Uytterhaegen et al., 1994). Degradation of structural proteins such as nebulin, titin and desmin could destroy the integrity of myofibrils (Taylor et al., 1995). However, cytoskeletal proteins such as titin, nebulin and desmin account for a small percentage of the myofibril, while the main body parts of the myofibril include the thin filament composed of actin and the thick filament composed of myosin (Clark et al., 2002). Our previous research found that the direct depolymerizing progress of actin filaments was positively correlated with myofibrillar fraction and tenderization (Zhou et al., 2016). In addition, we demonstrated that the disruption of actin filaments by calcium ion treatments could contribute to the tenderization of goose meat (Li et al., 2017). As an innovative technology, ultrasound treatments (UT) have been applied in the meat industry to promote curing rate, improve the rheological and structural properties of chicken myofibrillar gel and increase tenderness and water holding capacity of red meat during recent years (Kang et al., 2016; Kang et al., 2017; Wang et al., 2017). High-intensity UT (2002 kHz) has been used to improve the textural parameters and decrease the filtering residue of beef (Barekat and Soltanizadeh, 2016). Ultrasound increased the releases of calcium thus improve the tenderness (Turantaş et al., 2015). Lyng et al. (1998) reported that low-intensity UT (20 kHz) enhanced tenderness in lamb muscles. Kang et al. (2017) reported that UT at a frequency of 20 kHz improved the WHC and tenderness of beef by accelerating myofibrillar fraction and proteolysis of desmin and troponin-T. However, the effect of ultrasound treatment on the quality of poultry meat is not well understood. Got et al. (1999) demonstrated that UT could cause an ultrastructural alteration in the Z-line region. It is known that the Z-disk plays an important role in stabilizing actin filament during contraction (Sequeira et al., 2014). The disruption of Z-disk integrity by UT during conditioning is possible to cause the depolymerization of actin filament. Until now there has been limited studies on UT to improve the tenderization of goose meat by the depolymerization of F-actin. In order to clarify the possible tenderizing mechanisms of ultrasound by the depolymerization of actin filament, the present study was to evaluate the influence of UT on the change of shear force, cooking loss, myofibrillar fraction and particle size, the actin filaments morphology, and the equilibrium of G-actin and F-actin contents during conditioning. MATERIALS AND METHODS Treatments and Sampling Thirty-two Eastern Zhejiang White male Geese of 75-day-old with living weight 4, 205 ± 85 g were slaughtered by cutting the carotid artery and the jugular vein in the neck region from a local processing plant with no stress according to the permission of Ningbo University Animal Welfare Committee. The geese breast muscles were taken out of geese after 15 min of animal exsanguinations. 46 geese breast muscles were cut into 184 strips (20 × 10 × 10 mm for each) parallel to fiber direction (every breast muscle almost was cut in 4 strips); 18 geese breast muscles were cut into 72 cubes (0.8 cm3) parallel to fiber direction (every breast muscle almost was cut in 4 cubes). The 60 cubes and 180 strips with standard sizes, straight fiber and invisible connective tissue were selected from all of cubes and strips and then divided into three equal groups randomly. The 20 cubes and 60 strips were prepared for each group. Samples were put into a bag (oxygen, carbon dioxide and water vapor transmission rates of the film were 14.483 cm3/(m2 × 24 h × atm), 63.683 cm3/(m2 × 24 h × atm) and 54 g/(m2 × 24 h), respectively) to vacuumize with double-chamber vacuum packing machine (Yizhong machinery co., Ltd, Zhucheng, China) in the vacuum level of 200 pa. Three groups were treated with non-ultrasound, 300 W ultrasound and 600 W ultrasound respectively. One group was treated with an ultrasonic processor (KQ-300DE, Kunshan Ultrasonic Instrument Co., Ltd., China) by a power of 300 W at 40 kHz for 30 min as 300 W group. Another group was treated with an ultrasonic processor (KQ-600DE, Kunshan Ultrasonic Instrument Co., Ltd., China) by a power of 600 W at 40 kHz for 30 min as 600 W group. The rest non-ultrasound treated samples were considered as the control. At the end of the treatment, 5 cubes and 10 strips from each group were taken immediately as the defined samples at 0 h. The rest of cubes and strips were transferred to plastic trays. Then these cubes and strips were enclosed with a low-density polyethylene plastic film (oxygen, carbon dioxide and water vapor transmission rates of the film were 14, 483 cm3/(m2 × 24 h × atm), 63, 683 cm3/(m2 × 24 h × atm) and 54 g/(m2 × 24 h), respectively) and were stored at 4°C for 168 h. At each stored point (0, 8, 24, 48, 96 and 168 h), 10 strips from each group were taken immediately. The 3 strips (10 mm × 10 mm × 10 mm) for shear force determination and 3 strips for cooking loss determination were made from 10 strips in each of point parallel the fiber direction; the rest of 4 strips in each point in three groups were taken immediately, frozen in liquid nitrogen and stored at −80°C until being used for F-actin, G-actin, myofibril fraction index (MFI) and particle size determinations. The 5 cubes of each group at 0, 24, 96 and 168 h were also taken and frozen in liquid nitrogen for cryosections respectively and stored at −80°C until being analyzed. Shear Force Measurement Shear force measurement was done as described by Marino et al. (2013). Shear force samples were individually enclosed with retort pouch and cooked in a water bath at 85°C until the internal temperature of samples reached 75°C with a K-thermocouple (Shanghai Xinghua Instrument Factory, China). The peak force values (kg) were determined by placing the cooked samples perpendicular to the longitudinal axis of the muscle fibers under Warner-Bratzler shear blade on XL1155 equipment (Xielikeji Co. Ltd, Herbin, China). The temperature of the samples when sheared was 75°C. Each sample was sheared perpendicular to the fiber at 100 mm/min cross-head speed. The average peak force values of 5 measurements from each sampling were considered as shear force values. Cooking Loss Measurement Cooking loss was measured according to Klinhom et al. (2015). The samples were placed in retort pouch and cooked in a water bath at 80°C until the internal temperature of 70°C was reached with a K-thermocouple (Shanghai Xinghua Instrument Factory, China). Cooked samples were cooled to room temperature, blotted dry and reweighed. Cooking loss was calculated by the percentage of difference between different weights before and after cooking. The Measurement of MFI The measurement of MFI was done as described previously by Allane et al. (2010). Briefly, samples (0.5 g) avoiding any visible fat or connective tissue were taken and homogenized in 10 mL of ice cold buffer at 4°C with a DY89-I high speed homogenizer (Scientzco., Ningbo, China) for 3 × 10 s at 10, 000 rpm. After the sample was homogenized, the myofibrillar suspensions were filtered through mesh strainers (Culina, UK) with 1 mm2 holes into 50 mL centrifuge tubes to remove the connective tissue. The suspensions with 10 mL of cold buffer were filtrated through the mesh strainers with 1 mm2 holes again. The filtrates were centrifuged with a refrigerated centrifuge (Hunan Xiangyi Laboratory Instrument Development Co., Changsha, China) at 1, 000 g for 20 min at 4°C. The precipitation of myofibril was resuspended in a cold 10 mL buffer. Suspension was vortexed and centrifuged again. This process was repeated twice and the precipitation finally was resuspended in a cold 10 mL buffer. A bicinchoninic acid (BCA) protein assay kit (Thermo Scientific, MA) was used to determine the protein content of supernatant. The absorption was measured at 562 nm using with a 96-Well Plate Reader M200 (Tecan, Austria). A bovine serum albumin standard curve was used. Suspensions were diluted with buffer to a final protein concentration of 0.5 mg/mL. The diluted suspensions were poured into a cuvette and mixed. The absorbance measured was immediately at 540 nm using a 96-Well Plate Reader M200. The mean of five absorbance readings was multiplied by 150 which was the MFI. Particle Size Measurement The measurement was done as described by Lametsch et al. (2007). Sample (2.5 g) was homogenized in 30 mL cold buffer (100 mM KCl, 10 mM KH2PO4, 10 mM K2HPO4, 1.0 mM EDTA, 1.0 mM MgCl2) using a DY89-I high speed homogenizer (Scientzco., Ningbo, China) for 3 × 10 s at 10, 000 rpm at 4°C. The homogenate was centrifuged with a refrigerated centrifuge (Hunan Xiangyi Laboratory Instrument Development Co., Changsha, China) at 1, 000 g for 15 min at 4°C. The supernatant was discarded and the myofibrils were suspended in 25 mL buffer and centrifuged at 1, 000 g for 15 min at 4°C. The supernatant was discarded and the myofibrils were suspended in 15 mL buffer and subsequently filtered through mesh strainers (Culina, UK) with 1 mm2 holes to remove connective tissue and debris. The size distribution of the homogenate was measured using Mastersizer Micro (Malvern Instruments Ltd., Worcestershire, UK). The following indexes of proteins were analyzed using Mastersizer software (version 5.12c, Malvern Instruments Co. Ltd., Worcestershire, UK). D4,3 is the mean diameter in volume whereas D3,2 is the mean diameter in surface. Dv,0.1 is the size for which 10% of the sample particles have a lower size; Dv,0.5 is the size for which 50% of the sample particles have a lower size; Dv,0.9 is the particle size for which 90% of the sample particles have a lower size. Morphological Analysis for Actin Filaments The histology analysis was done as described by our previous study of Li et al. (2017). Each cube was cut into 8 μm sections with a Leica Frigicut cryostat (Leica, Nussloch, Germany). The slices were attached to Apes (Sigma, St. Louis) coated slides. The cryosections were fixed in 4% paraformaldehyde with 10 mM PBS (137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2HPO4, 1.4 mM KH2PO4, pH 7.4) at room temperature for 1 h. Then the cryosections were washed in 10 mM PBS for 15 min. Then they were permeabilized with 0.2% Triton X-100 (Beijing Sdarbio Science & Technology Co., Ltd, Beijing, China) in 10 mM PBS for 7 min and washed in 10 mM PBS for 15 min again. The cryosections were incubated in 0.1 μM FITC-phalloidin (Molecular Probes, cytoskeleton, Inc) for 1 h and washed in 10 mM PBS for 15 min. Actin filaments were observed by IX71 fluorescence microscope (Olympus, Japan). Extraction and Determination of F-Actin The F-actin content was determined as described previously by Zhou et al. (2016). Briefly, 0.5 g of samples were homogenized in 5 mL of lysis buffer by DY89-I high speed homogenizer (Scientz co., Ningbo, China) at 10, 000 rpm for 3 × 10 s while cooled in ice. The homogenate was incubated in 200 μL of 0.6 μM FITC-phalloidin at 4°C for 2 h. The samples were centrifuged at 80, 000 g for 60 min at 4°C in ultracentrifuge (Beckman Instruments, Inc., Palo Alto, CA). The pellet was extracted in 1 mL of methanol for 24 h. The fluorescence of extracted solution was continuously monitored at 540 nm as excitation wavelengths and at 575 nm as emission wavelengths by using a 96-Well Black Plate Reader M200 (Tecan, Austria) at room temperature. The intensity of fluorescence per-gram of muscle tissue was expressed as arbitrary unit (a.u.). Extraction of G-Actin The content of G-actin was determined as described originally by Cano et al. (1991). Briefly, 1.0 g of muscle samples were homogenized with 8 mL of lysis buffer by a DY89-I high speed homogenizer (Scientz co., Ningbo, China) for 3 × 10 s at 10, 000 rpm while cooled in ice. The homogenate was centrifuged at 6, 000 g for 20 min at 4°C with a refrigerated centrifuge (Hunan Xiangyi Laboratory Instrument Development Co., Changsha, China). Suspensions were diluted with lysis buffer to the same protein concentration (mg/mL) by using a bicinchoninic acid (BCA) protein assay kit (Thermo Scientific, MA). The final supernatants were used for G-actin assays. Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) and Western Blotting The SDS-PAGE and western blotting were done to determine the levels of G-actin as described previously by Warner et al. (1997). Glyceraldehyde-3-phosphate dehydrogenas (GAPDH), with the molecular weight of 36 kDa, was used as standard reference for Western blot. G-actin sample (20 μg) was run on a 12.5% SDS-PAGE and then transferred to Polyvinylidene fluoride membranes (Immobilon-P, Millipore, MA). Membranes were blocked in 1 × PBST containing 5% fat-free dry milk for 1 h at 25°C and incubated with primary antibody for 24 h at 4°C. The primary antibodies of rabbit anti-beta-actin (ab8227, 1:5000, Abcam, Cambridge, MA) and mouse anti-GAPDH (ab8245, 1:5000, Abcam, Cambridge, MA) were diluted in 1 × PBST solution containing 0.5% fat-free dry milk, respectively. The goat anti-rabbit HRP-conjugated secondary antibodies (ab97051, 1:10,000, Abcam, Cambridge, MA) and goat anti-mouse HRP-conjugated secondary antibodies (ab97023, 1:10,000, Abcam, Cambridge, MA) were diluted in 1 × PBST solution containing 0.5% fat-free dry milk, respectively. According to instructions, the validation of the antibodies were suitable for western blot. Membranes were incubated with secondary antibodies at 25°C for 1 h. ECL Plus Western blotting detection kit (Amersham Biosciences, UK) was used to detect the blot according to the manufacturer's instructions. Western blot data were quantified by specialized software and a microcomputer (Quantity One Software, Bio-Rad Laboratories, PA). Statistical Analysis All results were expressed as means of three replications. All data were presented as mean ± standard error. Statistical analysis was carried out by the multiple-factors (treatments and storage time, including two-way interactions) analysis of variance (ANOVA) procedure (Duncan's Multiple Range Test) of SAS 8.0 software (SAS Institute Inc., Cary, NC) to analyze the change and the difference of shear force, cooking loss, MFI, particle size, F-actin contents and expression levels of G-actin among three treatments (control, 300 W ultrasound and 600 W ultrasound) during 168 h of storage. The significant level was set as 0.05. RESULTS AND DISCUSSION The Changes of Shear Force in three Groups The changes of shear force are shown in Figure 1A in three groups. The values of shear force significantly decreased during 24–168 h in control group and decreased during 8–168 h in 300 W and 600 W group (P < 0.05). Among three groups, the values of shear force in 600 W group were significantly lower during 24–168 h than 300 W group and control group (P < 0.05). The values of shear force in 300 W group were significantly lower at 24, 96 and 168 h than control group (P < 0.05). Our research indicated that UT reduced the shear force of goose meat. Figure 1. View largeDownload slide The changes of shear force (A) and cooking loss (B) in three groups during conditioning. a-i Identical letters indicated that there were no significant difference in different treatments (P > 0.05). Figure 1. View largeDownload slide The changes of shear force (A) and cooking loss (B) in three groups during conditioning. a-i Identical letters indicated that there were no significant difference in different treatments (P > 0.05). Consumer tenderness acceptability increased as shear force values decreased (Miller et al., 2001). Smith et al. (2002) observed that shear force values of bovine Semitendinosus muscle reduced due to ultrasonic treatment. It was reported that the cavitation generated by UT causes the physical disruption of meat (Alarconrojo et al., 2015). UT has been found to improve tenderness by the disruption of myofibrillar integrity (Got et al., 1999). The Change of Cooking Loss in three Groups The changes of cooking loss are shown in Figure 1B in three groups. The values of cooking loss significantly decreased during 24–168 h in control group, decreased during 8–168 h in 300 W group and decreased during 8–168 h in 600 W group (P < 0.05). Among three groups, the values of cooking loss in 600 W group were significantly lower during 24–168 h than 300 W group and control group (P < 0.05). The values of cooking loss in 300 W group were lower during 48–168 h than control group (P < 0.05). Our study indicated that UT improved the water-holding capacity of goose meat. Dolatowski et al. (2000) reported that UT changed the structure of meat tissue (especially protein molecules and cell) and increased the water-holding capacity of meat. Vimini et al. (2010) stated that UT could reduce cooking loss of meat, which was consistent with our results. It has been found that UT increased the water binding strength of meat which is responsible for improving cooking loss (Reynolds et al., 2010). Alarconrojo et al. (2015) assumed that ultrasound reduced cooking losses of meat by weakening connective tissues and myofibrillar structure. The Changes of MFI and Particle size in Three Groups The changes of MFI are shown in Figure 2 in three groups. The values of MFI significantly increased during 24–168 h in 300 W group and control group, during 8–168 h in 600 W group (P < 0.05). Among three treatments, samples in 600 W group showed the higher values of MFI at 48, 96 and 168 h than in 300 W group and control group (P < 0.05). The MFI values of 300 W group were significantly higher at 48, 96 and 168 h than control group (P < 0.05). Figure 2. View largeDownload slide The changes of myofibrillar fragmentation index in three groups during conditioning. a-j Identical letters indicated that there were no significant difference in different treatments (P > 0.05). Figure 2. View largeDownload slide The changes of myofibrillar fragmentation index in three groups during conditioning. a-j Identical letters indicated that there were no significant difference in different treatments (P > 0.05). Changes in D4,3, D3,2, Dv,0.1, Dv,0.5, Dv,0.9 of particle size in three groups during conditioning are shown in Table 1. The values of D4,3, D3,2, Dv,0.1 and Dv,0.9 significantly decreased during 24–168 h in the control while the value of Dv,0.5 in the control significantly decreased during 8–168 h (P < 0.05). The values of D4,3, D3,2, Dv,0.1 and Dv,0.5 significantly decreased during 24–168 h in the 300 W group, while the value of Dv,0.9 in the 300 W group significantly decreased during 8–168 h (P < 0.05). The values of D4,3, Dv,0.1, Dv,0.5 and Dv,0.9 significantly decreased during 8–168 h in the 600 W group, while the value of D3,2 in the 600 W group significantly decreased during 24–168 h (P < 0.05). Among three groups, the values of D4,3, D3,2, Dv,0.1 and Dv,0.5 in 600 W group were lower during 48–168 h than 300 W group and control group, while the value of Dv,0.9 in 600 W group were significantly lower during 24–168 h than 300 W group and control group (P < 0.05). The values of D3,2, Dv,0.1 and Dv,0.5 in 300 W group were significantly lower during 48–168 h than the control and the value of D4,3 in 300 W group were significantly lower during 48–96 h than the control, while the value of Dv,0.9 in 300 W group were significantly lower during 96–168 h than control group (P < 0.05). The particle size distributions showed that UT increased the level of smaller particles and decreased the level of large particles compared to the control. Table 1. Changes in average particle size of myofibrillar proteins in three groups during conditioning.   D4,3/μm  D3,2/μm  DV,0.1/μm  DV,0.5/μm  DV,0.9/μm  CK-0h  196.55 ± 1.48a  61.7 ± 1.33a  32.83 ± 1.12a  140.36 ± 1.14a  423.25 ± 2.98a  CK-8h  195.27 ± 1.17a,b  59.38 ± 1.23a,b  31.84 ± 1.26a,b  138.31 ± 0.76a  418.91 ± 2.39a,b  CK-24h  193.41 ± 1.28b  57.27 ± 1.33b,c  30.45 ± 1.28b  135.68 ± 1.32b  412.53 ± 2.34b,c  CK-48h  188.01 ± 1.2d  52.2 ± 1.75d  28.67 ± 0.96c  129.46 ± 0.88c  398.57 ± 3.99d,e  CK-96h  183.45 ± 1.11f  47.43 ± 1.38e,f  25.42 ± 1.15d  124.02 ± 1.17d,e  385.38 ± 3.34e,f  CK-168h  179.33 ± 1.02 g,h  43.39 ± 1.35g  22.6 ± 1.14f  116.5 ± 0.89f  372.34 ± 2.88h  300W-0h  196 ± 0.54a  60.71 ± 1.4a,b  32.42 ± 0.6a,b  139.22 ± 0.91a  421.5 ± 2.38a,b  300W-8h  194.89 ± 0.37a,b  58.42 ± 1.18b  31.21 ± 0.65a,b  137.38 ± 0.73a,b  414.71 ± 2.26b  300W-24h  192.67 ± 0.8b,c  56.45 ± 1.35b,c  29.63 ± 1.27b,c  132.4 ± 1.11b,c  407.48 ± 2.59c  300W-48h  185.78 ± 1.09e  49.46 ± 1.26e  26.24 ± 0.79d  126.69 ± 0.83d  391 ± 2.46e  300W-96h  180.79 ± 1.2g  43.22 ± 1.12g  23.93 ± 0.91e  116.5 ± 1.09f  378.64 ± 2.49g  300W-168h  176.5 ± 1.38h  40.14 ± 1.14h  20.6 ± 0.98g  110.72 ± 1.15g  365.63 ± 1.2i  600W-0h  195.3 ± 0.82a,b  60.45 ± 1.23a,b  31.93 ± 0.85a,b  138.45 ± 0.77a  418.91 ± 1.09a,b  600W-8h  193.7 ± 0.79b  58.14 ± 1.09b,c  30.4 ± 1.07b  136.54 ± 1.31a,b  412.67 ± 1.64b,c  600W-24h  190.7 ± 0.72c  55.47 ± 1.5c  28.33 ± 1.04c  130.21 ± 1.24c  400.96 ± 1.75d  600W-48h  181.67 ± 0.88g  47.43 ± 1.06f  24.73 ± 1e  123.22 ± 0.89e  383.64 ± 2.06f  600W-96h  177.19 ± 0.83h  41.25 ± 1.11h  22.42 ± 0.96f  111.59 ± 1.48g  371.45 ± 1.54h  600W-168h  174.45 ± 0.92i  38.15 ± 1.34i  19.43 ± 0.9h  105.63 ± 1.58h  359.57 ± 2.45j    D4,3/μm  D3,2/μm  DV,0.1/μm  DV,0.5/μm  DV,0.9/μm  CK-0h  196.55 ± 1.48a  61.7 ± 1.33a  32.83 ± 1.12a  140.36 ± 1.14a  423.25 ± 2.98a  CK-8h  195.27 ± 1.17a,b  59.38 ± 1.23a,b  31.84 ± 1.26a,b  138.31 ± 0.76a  418.91 ± 2.39a,b  CK-24h  193.41 ± 1.28b  57.27 ± 1.33b,c  30.45 ± 1.28b  135.68 ± 1.32b  412.53 ± 2.34b,c  CK-48h  188.01 ± 1.2d  52.2 ± 1.75d  28.67 ± 0.96c  129.46 ± 0.88c  398.57 ± 3.99d,e  CK-96h  183.45 ± 1.11f  47.43 ± 1.38e,f  25.42 ± 1.15d  124.02 ± 1.17d,e  385.38 ± 3.34e,f  CK-168h  179.33 ± 1.02 g,h  43.39 ± 1.35g  22.6 ± 1.14f  116.5 ± 0.89f  372.34 ± 2.88h  300W-0h  196 ± 0.54a  60.71 ± 1.4a,b  32.42 ± 0.6a,b  139.22 ± 0.91a  421.5 ± 2.38a,b  300W-8h  194.89 ± 0.37a,b  58.42 ± 1.18b  31.21 ± 0.65a,b  137.38 ± 0.73a,b  414.71 ± 2.26b  300W-24h  192.67 ± 0.8b,c  56.45 ± 1.35b,c  29.63 ± 1.27b,c  132.4 ± 1.11b,c  407.48 ± 2.59c  300W-48h  185.78 ± 1.09e  49.46 ± 1.26e  26.24 ± 0.79d  126.69 ± 0.83d  391 ± 2.46e  300W-96h  180.79 ± 1.2g  43.22 ± 1.12g  23.93 ± 0.91e  116.5 ± 1.09f  378.64 ± 2.49g  300W-168h  176.5 ± 1.38h  40.14 ± 1.14h  20.6 ± 0.98g  110.72 ± 1.15g  365.63 ± 1.2i  600W-0h  195.3 ± 0.82a,b  60.45 ± 1.23a,b  31.93 ± 0.85a,b  138.45 ± 0.77a  418.91 ± 1.09a,b  600W-8h  193.7 ± 0.79b  58.14 ± 1.09b,c  30.4 ± 1.07b  136.54 ± 1.31a,b  412.67 ± 1.64b,c  600W-24h  190.7 ± 0.72c  55.47 ± 1.5c  28.33 ± 1.04c  130.21 ± 1.24c  400.96 ± 1.75d  600W-48h  181.67 ± 0.88g  47.43 ± 1.06f  24.73 ± 1e  123.22 ± 0.89e  383.64 ± 2.06f  600W-96h  177.19 ± 0.83h  41.25 ± 1.11h  22.42 ± 0.96f  111.59 ± 1.48g  371.45 ± 1.54h  600W-168h  174.45 ± 0.92i  38.15 ± 1.34i  19.43 ± 0.9h  105.63 ± 1.58h  359.57 ± 2.45j  a-jIdentical letters indicate that there is no significant difference in different processing points (P > 0.05). View Large Table 1. Changes in average particle size of myofibrillar proteins in three groups during conditioning.   D4,3/μm  D3,2/μm  DV,0.1/μm  DV,0.5/μm  DV,0.9/μm  CK-0h  196.55 ± 1.48a  61.7 ± 1.33a  32.83 ± 1.12a  140.36 ± 1.14a  423.25 ± 2.98a  CK-8h  195.27 ± 1.17a,b  59.38 ± 1.23a,b  31.84 ± 1.26a,b  138.31 ± 0.76a  418.91 ± 2.39a,b  CK-24h  193.41 ± 1.28b  57.27 ± 1.33b,c  30.45 ± 1.28b  135.68 ± 1.32b  412.53 ± 2.34b,c  CK-48h  188.01 ± 1.2d  52.2 ± 1.75d  28.67 ± 0.96c  129.46 ± 0.88c  398.57 ± 3.99d,e  CK-96h  183.45 ± 1.11f  47.43 ± 1.38e,f  25.42 ± 1.15d  124.02 ± 1.17d,e  385.38 ± 3.34e,f  CK-168h  179.33 ± 1.02 g,h  43.39 ± 1.35g  22.6 ± 1.14f  116.5 ± 0.89f  372.34 ± 2.88h  300W-0h  196 ± 0.54a  60.71 ± 1.4a,b  32.42 ± 0.6a,b  139.22 ± 0.91a  421.5 ± 2.38a,b  300W-8h  194.89 ± 0.37a,b  58.42 ± 1.18b  31.21 ± 0.65a,b  137.38 ± 0.73a,b  414.71 ± 2.26b  300W-24h  192.67 ± 0.8b,c  56.45 ± 1.35b,c  29.63 ± 1.27b,c  132.4 ± 1.11b,c  407.48 ± 2.59c  300W-48h  185.78 ± 1.09e  49.46 ± 1.26e  26.24 ± 0.79d  126.69 ± 0.83d  391 ± 2.46e  300W-96h  180.79 ± 1.2g  43.22 ± 1.12g  23.93 ± 0.91e  116.5 ± 1.09f  378.64 ± 2.49g  300W-168h  176.5 ± 1.38h  40.14 ± 1.14h  20.6 ± 0.98g  110.72 ± 1.15g  365.63 ± 1.2i  600W-0h  195.3 ± 0.82a,b  60.45 ± 1.23a,b  31.93 ± 0.85a,b  138.45 ± 0.77a  418.91 ± 1.09a,b  600W-8h  193.7 ± 0.79b  58.14 ± 1.09b,c  30.4 ± 1.07b  136.54 ± 1.31a,b  412.67 ± 1.64b,c  600W-24h  190.7 ± 0.72c  55.47 ± 1.5c  28.33 ± 1.04c  130.21 ± 1.24c  400.96 ± 1.75d  600W-48h  181.67 ± 0.88g  47.43 ± 1.06f  24.73 ± 1e  123.22 ± 0.89e  383.64 ± 2.06f  600W-96h  177.19 ± 0.83h  41.25 ± 1.11h  22.42 ± 0.96f  111.59 ± 1.48g  371.45 ± 1.54h  600W-168h  174.45 ± 0.92i  38.15 ± 1.34i  19.43 ± 0.9h  105.63 ± 1.58h  359.57 ± 2.45j    D4,3/μm  D3,2/μm  DV,0.1/μm  DV,0.5/μm  DV,0.9/μm  CK-0h  196.55 ± 1.48a  61.7 ± 1.33a  32.83 ± 1.12a  140.36 ± 1.14a  423.25 ± 2.98a  CK-8h  195.27 ± 1.17a,b  59.38 ± 1.23a,b  31.84 ± 1.26a,b  138.31 ± 0.76a  418.91 ± 2.39a,b  CK-24h  193.41 ± 1.28b  57.27 ± 1.33b,c  30.45 ± 1.28b  135.68 ± 1.32b  412.53 ± 2.34b,c  CK-48h  188.01 ± 1.2d  52.2 ± 1.75d  28.67 ± 0.96c  129.46 ± 0.88c  398.57 ± 3.99d,e  CK-96h  183.45 ± 1.11f  47.43 ± 1.38e,f  25.42 ± 1.15d  124.02 ± 1.17d,e  385.38 ± 3.34e,f  CK-168h  179.33 ± 1.02 g,h  43.39 ± 1.35g  22.6 ± 1.14f  116.5 ± 0.89f  372.34 ± 2.88h  300W-0h  196 ± 0.54a  60.71 ± 1.4a,b  32.42 ± 0.6a,b  139.22 ± 0.91a  421.5 ± 2.38a,b  300W-8h  194.89 ± 0.37a,b  58.42 ± 1.18b  31.21 ± 0.65a,b  137.38 ± 0.73a,b  414.71 ± 2.26b  300W-24h  192.67 ± 0.8b,c  56.45 ± 1.35b,c  29.63 ± 1.27b,c  132.4 ± 1.11b,c  407.48 ± 2.59c  300W-48h  185.78 ± 1.09e  49.46 ± 1.26e  26.24 ± 0.79d  126.69 ± 0.83d  391 ± 2.46e  300W-96h  180.79 ± 1.2g  43.22 ± 1.12g  23.93 ± 0.91e  116.5 ± 1.09f  378.64 ± 2.49g  300W-168h  176.5 ± 1.38h  40.14 ± 1.14h  20.6 ± 0.98g  110.72 ± 1.15g  365.63 ± 1.2i  600W-0h  195.3 ± 0.82a,b  60.45 ± 1.23a,b  31.93 ± 0.85a,b  138.45 ± 0.77a  418.91 ± 1.09a,b  600W-8h  193.7 ± 0.79b  58.14 ± 1.09b,c  30.4 ± 1.07b  136.54 ± 1.31a,b  412.67 ± 1.64b,c  600W-24h  190.7 ± 0.72c  55.47 ± 1.5c  28.33 ± 1.04c  130.21 ± 1.24c  400.96 ± 1.75d  600W-48h  181.67 ± 0.88g  47.43 ± 1.06f  24.73 ± 1e  123.22 ± 0.89e  383.64 ± 2.06f  600W-96h  177.19 ± 0.83h  41.25 ± 1.11h  22.42 ± 0.96f  111.59 ± 1.48g  371.45 ± 1.54h  600W-168h  174.45 ± 0.92i  38.15 ± 1.34i  19.43 ± 0.9h  105.63 ± 1.58h  359.57 ± 2.45j  a-jIdentical letters indicate that there is no significant difference in different processing points (P > 0.05). View Large MFI is one of the most widely useful indicator to determine postmortem proteolysis and tenderization in meat (Volpelli et al., 2005). Kang et al. (2017) reported that beef treated with ultrasound could increase MFI values thus increase tenderness of meat. Karumendu et al. (2009) demonstrated that the increase in MFI is accompanied by a decrease in shear force. In our study, the increase of MFI by UT contributed to the decrease of Warner-Bratzler shear force values and cooking loss during conditioning. Shin et al. (2008) indicated that the decrease of particle size of myofibril proteins contributed to meat tenderization. Li et al. (2014) reported that UT (450 W) caused a decrease in particle size of myofibril fragmentation in chicken breast meat since the distribution became more uniform and narrowed. The reduction of particle size is due to proteins dissociation during cavitation by UT and particles under sonication were violently agitated which resulted in broken aggregate particles (Zhang et al., 2017). It demonstrated that the reduction of particle size for myofibril fragmentation was contributed to the decrease of shear force values at storage periods directly (Crouse et al., 1991). The smaller particle size contributed to stronger protein-water interactions which strengthened proteins solubility and water holding capacity (Jambrak et al., 2014). The decrease in particle size by UT was due to the disruption of myofibril structure (Jayasooriya et al., 2004). In our research, the decrease in particle size by UT was related to the result of the increase in MFI. The Morphological Change of Actin Filaments in Three Groups The changes of actin filaments by FITC-phalloidin staining in three groups are shown in Figure 3. Actin filaments morphology did not show significant fracture among three treatments at 0 h. In the control, actin filaments disrupted slightly during 96–168 h. Actin filaments structure in 300 W group changed more dramatically during 96–168 h than the control. Actin filaments in 600 W group disrupted more dramatically during 96–168 h than 300 W group and the control. It indicated that ultrasound treatment accelerated the proteolysis of actin filaments. Figure 3. View largeDownload slide The changes of actin filaments in three groups during conditioning. CK, 300 W and 600 W indicated control, 300 W ultrasound group and 600 W ultrasound group, respectively. Images were obtained using a fluorescence microscope with the 100 × objective (Olympus, Japan). Arrows in 300 W and 600 W groups implied that the actin filaments were disrupted dramatically. Figure 3. View largeDownload slide The changes of actin filaments in three groups during conditioning. CK, 300 W and 600 W indicated control, 300 W ultrasound group and 600 W ultrasound group, respectively. Images were obtained using a fluorescence microscope with the 100 × objective (Olympus, Japan). Arrows in 300 W and 600 W groups implied that the actin filaments were disrupted dramatically. FITC-phalloidin enabled us to observe the distribution of F-actin in myofibril structure (Schmit and Lambert, 1990). The basic contractile unit of myofibrils is the sarcomere which include myosin, actin and nebulin filament (Littlefield and Fowler, 2008). Meat treated with ultrasound resulted in the disintegration of the myofibrillar scaffold (Chen et al., 2015). Ahn et al. (2003) thought that the increase of MFI with aging may be related to the breaking of myofibrillar proteins into segments near the Z-disk. Latoch (2010) indicated that UT can induce the degradation of I-Z-I band proteins and then increase the MFI. It was noted that actin accounts for 22% of myofibrillar protein and that its content was higher than the level of some cytoskeletal proteins in Z-disk substantially (Yates and Greaser, 1983). We thought that the degradation of cytoskeletal proteins near the Z-disk was not the directly reason for the increase of MFI. Our result indicated that the disruption of actin filaments but not Z-disks by UT could directly lead to the increase of MFI and the decrease of particle size. It could be an important factor in meat tenderization during conditioning. Changes of F-Actin and G-Actin Contents in Three Treatments The changes of F-actin content in myofibrils are shown in Figure 4. The content of F-actin in all treatments had the highest values at 0 h which indicated a very low extent of F-actin depolymerization. The content of F-actin significantly reduced during 8–168 h in the control and 300 W group and decreased during 0–168 h in 300 W group (P < 0.05). 600 W group showed lower content of F-actin during 8–168 h compared to the control and 300 W group (P < 0.05). Compared to control group, the content of F-actin was significantly lower during 24–168 h in 300 W group (P < 0.05). Figure 4. View largeDownload slide The changes of F-actin content in myofibrils in three groups during conditioning. a-j Identical letters indicated that there were no significant difference in different treatments (P > 0.05). Figure 4. View largeDownload slide The changes of F-actin content in myofibrils in three groups during conditioning. a-j Identical letters indicated that there were no significant difference in different treatments (P > 0.05). The changes of G-actin (43 kDa) levels are shown in Figure 5 by western blot analysis in three groups. The G-actin bands significantly increased during 0–24 h and during 96–168 h in control group, during 24–96 h in 300 W group and 600 W group (P < 0.05), respectively. The G-actin bands in 600 W group had larger values from 24 to 96 h than in the control and 300 W group (P < 0.01). The G-actin bands of 300 W group had higher values during 96–168 h than the control (P < 0.05). The decrease of F-actin and increase of G-actin implied that F-actin depolymerized to G-actin. The rate of the depolymerization of F-actin to G-actin in 600 W group was faster than 300 W group; the rate in the control was the slowest among three groups. Figure 5. View largeDownload slide The changes of G-actin levels by SDS-PAGE and western blot in three groups during conditioning. a-e Identical letters in the line indicated that there were no significant difference in different treatments (P > 0.05). Figure 5. View largeDownload slide The changes of G-actin levels by SDS-PAGE and western blot in three groups during conditioning. a-e Identical letters in the line indicated that there were no significant difference in different treatments (P > 0.05). The phallacidin-stained F-actin of tissues and cells is widely used to quantify the content of F-actin (Sampath and Pollard, 1991). UT lead to a decrease in the expression of cytoskeletal proteins such as F-actin by immunocytochemical staining (Monici et al., 2007). Saleem et al. (2015) reported that ultrasonication causes the structural changes in myosin and actin of chicken breast muscle. It has been shown that nebulin participates in the stabilization of F-actin (Wang and Wright, 1988). Xiong et al. (2012) demonstrated that nebulin were degraded from chicken breast muscle by UT. It indicated that ultrasound could weaken F-actin by degraded nebulin proteins. Littlefield et al. (2001) indicated that there is an actin dynamics fluctuating between actin filaments and monomers in skeletal muscle cells. It is known that treatment with ultrasound (sonication) reinforce actin depolymerization (Carlier et al., 1984). UT could release Ca2+ which may regulated the pathway of the actin depolymerize to G-actin (Lange and Brandt, 1996). G-actin can be prepared from F-actin by the treatment of ultrasonic (Mihashi, 1964). A slow actin depolymerization began to occur after sonication, indicating that its alteration is caused by ultrasound extraction (Wendel and Dancker, 1986). In our results, we found that ultrasound treatments accelerated the transformation from F-actin to G-actin. CONCLUSIONS The disruption of actin filaments but not Z-disks by UT could contribute to myofibrillar fraction directly and the tenderization of goose meat. UT accelerated the transformation from F-actin to G-actin. 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The effect of ultrasound treatments on the tenderizing pathway of goose meat during conditioning

Poultry Science , Volume Advance Article – Apr 20, 2018

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Oxford University Press
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© 2018 Poultry Science Association Inc.
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0032-5791
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1525-3171
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10.3382/ps/pey143
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Abstract

ABSTRACT In order to figure out the effect of ultrasound treatments (UT) on the tenderizing pathway of goose meat, breast muscles of 32 Eastern Zhejiang White Geese were treated with different ultrasound powers (the control, 300 and 600 W) at 40 kHz for 30 min. Shear force, cooking loss, myofibrillar fraction index (MFI), particle size, morphological analysis of actin filaments and the levels of F-actin and G-actin were investigated during 168 h storage. Results showed that 600 W group had the lowest shear force and cooking loss at 24, 48, 96 and 168 h among treatments, while 300 W UT decreased shear force and cooking loss compared to the control. UT increased MFI and induced myofibrillar small particles (D3,2) by disrupting actin filaments for myofibril and transforming of F-actin to G-actin compared to the control. We concluded that UT tenderized goose meat by fragmenting actin filaments and myofibrillar fraction. INTRODUCTION Tenderness is one of the most important quality parameters affecting consumer satisfaction and the positive perception of meat products (Miller et al., 2001). It can be improved by various physical, chemical and biochemical methods, such as electric stimulation, high-pressure shockwaves, tumbling, calcium marination and plant proteases (Jung et al., 2000; Hwang et al., 2003; Alvarado and Sams, 2004; Naveena et al., 2004; Feng et al., 2017; Li et al., 2017). It was demonstrated that the mechanism of tenderization is related to the destruction of myofibrils (Uytterhaegen et al., 1994). Degradation of structural proteins such as nebulin, titin and desmin could destroy the integrity of myofibrils (Taylor et al., 1995). However, cytoskeletal proteins such as titin, nebulin and desmin account for a small percentage of the myofibril, while the main body parts of the myofibril include the thin filament composed of actin and the thick filament composed of myosin (Clark et al., 2002). Our previous research found that the direct depolymerizing progress of actin filaments was positively correlated with myofibrillar fraction and tenderization (Zhou et al., 2016). In addition, we demonstrated that the disruption of actin filaments by calcium ion treatments could contribute to the tenderization of goose meat (Li et al., 2017). As an innovative technology, ultrasound treatments (UT) have been applied in the meat industry to promote curing rate, improve the rheological and structural properties of chicken myofibrillar gel and increase tenderness and water holding capacity of red meat during recent years (Kang et al., 2016; Kang et al., 2017; Wang et al., 2017). High-intensity UT (2002 kHz) has been used to improve the textural parameters and decrease the filtering residue of beef (Barekat and Soltanizadeh, 2016). Ultrasound increased the releases of calcium thus improve the tenderness (Turantaş et al., 2015). Lyng et al. (1998) reported that low-intensity UT (20 kHz) enhanced tenderness in lamb muscles. Kang et al. (2017) reported that UT at a frequency of 20 kHz improved the WHC and tenderness of beef by accelerating myofibrillar fraction and proteolysis of desmin and troponin-T. However, the effect of ultrasound treatment on the quality of poultry meat is not well understood. Got et al. (1999) demonstrated that UT could cause an ultrastructural alteration in the Z-line region. It is known that the Z-disk plays an important role in stabilizing actin filament during contraction (Sequeira et al., 2014). The disruption of Z-disk integrity by UT during conditioning is possible to cause the depolymerization of actin filament. Until now there has been limited studies on UT to improve the tenderization of goose meat by the depolymerization of F-actin. In order to clarify the possible tenderizing mechanisms of ultrasound by the depolymerization of actin filament, the present study was to evaluate the influence of UT on the change of shear force, cooking loss, myofibrillar fraction and particle size, the actin filaments morphology, and the equilibrium of G-actin and F-actin contents during conditioning. MATERIALS AND METHODS Treatments and Sampling Thirty-two Eastern Zhejiang White male Geese of 75-day-old with living weight 4, 205 ± 85 g were slaughtered by cutting the carotid artery and the jugular vein in the neck region from a local processing plant with no stress according to the permission of Ningbo University Animal Welfare Committee. The geese breast muscles were taken out of geese after 15 min of animal exsanguinations. 46 geese breast muscles were cut into 184 strips (20 × 10 × 10 mm for each) parallel to fiber direction (every breast muscle almost was cut in 4 strips); 18 geese breast muscles were cut into 72 cubes (0.8 cm3) parallel to fiber direction (every breast muscle almost was cut in 4 cubes). The 60 cubes and 180 strips with standard sizes, straight fiber and invisible connective tissue were selected from all of cubes and strips and then divided into three equal groups randomly. The 20 cubes and 60 strips were prepared for each group. Samples were put into a bag (oxygen, carbon dioxide and water vapor transmission rates of the film were 14.483 cm3/(m2 × 24 h × atm), 63.683 cm3/(m2 × 24 h × atm) and 54 g/(m2 × 24 h), respectively) to vacuumize with double-chamber vacuum packing machine (Yizhong machinery co., Ltd, Zhucheng, China) in the vacuum level of 200 pa. Three groups were treated with non-ultrasound, 300 W ultrasound and 600 W ultrasound respectively. One group was treated with an ultrasonic processor (KQ-300DE, Kunshan Ultrasonic Instrument Co., Ltd., China) by a power of 300 W at 40 kHz for 30 min as 300 W group. Another group was treated with an ultrasonic processor (KQ-600DE, Kunshan Ultrasonic Instrument Co., Ltd., China) by a power of 600 W at 40 kHz for 30 min as 600 W group. The rest non-ultrasound treated samples were considered as the control. At the end of the treatment, 5 cubes and 10 strips from each group were taken immediately as the defined samples at 0 h. The rest of cubes and strips were transferred to plastic trays. Then these cubes and strips were enclosed with a low-density polyethylene plastic film (oxygen, carbon dioxide and water vapor transmission rates of the film were 14, 483 cm3/(m2 × 24 h × atm), 63, 683 cm3/(m2 × 24 h × atm) and 54 g/(m2 × 24 h), respectively) and were stored at 4°C for 168 h. At each stored point (0, 8, 24, 48, 96 and 168 h), 10 strips from each group were taken immediately. The 3 strips (10 mm × 10 mm × 10 mm) for shear force determination and 3 strips for cooking loss determination were made from 10 strips in each of point parallel the fiber direction; the rest of 4 strips in each point in three groups were taken immediately, frozen in liquid nitrogen and stored at −80°C until being used for F-actin, G-actin, myofibril fraction index (MFI) and particle size determinations. The 5 cubes of each group at 0, 24, 96 and 168 h were also taken and frozen in liquid nitrogen for cryosections respectively and stored at −80°C until being analyzed. Shear Force Measurement Shear force measurement was done as described by Marino et al. (2013). Shear force samples were individually enclosed with retort pouch and cooked in a water bath at 85°C until the internal temperature of samples reached 75°C with a K-thermocouple (Shanghai Xinghua Instrument Factory, China). The peak force values (kg) were determined by placing the cooked samples perpendicular to the longitudinal axis of the muscle fibers under Warner-Bratzler shear blade on XL1155 equipment (Xielikeji Co. Ltd, Herbin, China). The temperature of the samples when sheared was 75°C. Each sample was sheared perpendicular to the fiber at 100 mm/min cross-head speed. The average peak force values of 5 measurements from each sampling were considered as shear force values. Cooking Loss Measurement Cooking loss was measured according to Klinhom et al. (2015). The samples were placed in retort pouch and cooked in a water bath at 80°C until the internal temperature of 70°C was reached with a K-thermocouple (Shanghai Xinghua Instrument Factory, China). Cooked samples were cooled to room temperature, blotted dry and reweighed. Cooking loss was calculated by the percentage of difference between different weights before and after cooking. The Measurement of MFI The measurement of MFI was done as described previously by Allane et al. (2010). Briefly, samples (0.5 g) avoiding any visible fat or connective tissue were taken and homogenized in 10 mL of ice cold buffer at 4°C with a DY89-I high speed homogenizer (Scientzco., Ningbo, China) for 3 × 10 s at 10, 000 rpm. After the sample was homogenized, the myofibrillar suspensions were filtered through mesh strainers (Culina, UK) with 1 mm2 holes into 50 mL centrifuge tubes to remove the connective tissue. The suspensions with 10 mL of cold buffer were filtrated through the mesh strainers with 1 mm2 holes again. The filtrates were centrifuged with a refrigerated centrifuge (Hunan Xiangyi Laboratory Instrument Development Co., Changsha, China) at 1, 000 g for 20 min at 4°C. The precipitation of myofibril was resuspended in a cold 10 mL buffer. Suspension was vortexed and centrifuged again. This process was repeated twice and the precipitation finally was resuspended in a cold 10 mL buffer. A bicinchoninic acid (BCA) protein assay kit (Thermo Scientific, MA) was used to determine the protein content of supernatant. The absorption was measured at 562 nm using with a 96-Well Plate Reader M200 (Tecan, Austria). A bovine serum albumin standard curve was used. Suspensions were diluted with buffer to a final protein concentration of 0.5 mg/mL. The diluted suspensions were poured into a cuvette and mixed. The absorbance measured was immediately at 540 nm using a 96-Well Plate Reader M200. The mean of five absorbance readings was multiplied by 150 which was the MFI. Particle Size Measurement The measurement was done as described by Lametsch et al. (2007). Sample (2.5 g) was homogenized in 30 mL cold buffer (100 mM KCl, 10 mM KH2PO4, 10 mM K2HPO4, 1.0 mM EDTA, 1.0 mM MgCl2) using a DY89-I high speed homogenizer (Scientzco., Ningbo, China) for 3 × 10 s at 10, 000 rpm at 4°C. The homogenate was centrifuged with a refrigerated centrifuge (Hunan Xiangyi Laboratory Instrument Development Co., Changsha, China) at 1, 000 g for 15 min at 4°C. The supernatant was discarded and the myofibrils were suspended in 25 mL buffer and centrifuged at 1, 000 g for 15 min at 4°C. The supernatant was discarded and the myofibrils were suspended in 15 mL buffer and subsequently filtered through mesh strainers (Culina, UK) with 1 mm2 holes to remove connective tissue and debris. The size distribution of the homogenate was measured using Mastersizer Micro (Malvern Instruments Ltd., Worcestershire, UK). The following indexes of proteins were analyzed using Mastersizer software (version 5.12c, Malvern Instruments Co. Ltd., Worcestershire, UK). D4,3 is the mean diameter in volume whereas D3,2 is the mean diameter in surface. Dv,0.1 is the size for which 10% of the sample particles have a lower size; Dv,0.5 is the size for which 50% of the sample particles have a lower size; Dv,0.9 is the particle size for which 90% of the sample particles have a lower size. Morphological Analysis for Actin Filaments The histology analysis was done as described by our previous study of Li et al. (2017). Each cube was cut into 8 μm sections with a Leica Frigicut cryostat (Leica, Nussloch, Germany). The slices were attached to Apes (Sigma, St. Louis) coated slides. The cryosections were fixed in 4% paraformaldehyde with 10 mM PBS (137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2HPO4, 1.4 mM KH2PO4, pH 7.4) at room temperature for 1 h. Then the cryosections were washed in 10 mM PBS for 15 min. Then they were permeabilized with 0.2% Triton X-100 (Beijing Sdarbio Science & Technology Co., Ltd, Beijing, China) in 10 mM PBS for 7 min and washed in 10 mM PBS for 15 min again. The cryosections were incubated in 0.1 μM FITC-phalloidin (Molecular Probes, cytoskeleton, Inc) for 1 h and washed in 10 mM PBS for 15 min. Actin filaments were observed by IX71 fluorescence microscope (Olympus, Japan). Extraction and Determination of F-Actin The F-actin content was determined as described previously by Zhou et al. (2016). Briefly, 0.5 g of samples were homogenized in 5 mL of lysis buffer by DY89-I high speed homogenizer (Scientz co., Ningbo, China) at 10, 000 rpm for 3 × 10 s while cooled in ice. The homogenate was incubated in 200 μL of 0.6 μM FITC-phalloidin at 4°C for 2 h. The samples were centrifuged at 80, 000 g for 60 min at 4°C in ultracentrifuge (Beckman Instruments, Inc., Palo Alto, CA). The pellet was extracted in 1 mL of methanol for 24 h. The fluorescence of extracted solution was continuously monitored at 540 nm as excitation wavelengths and at 575 nm as emission wavelengths by using a 96-Well Black Plate Reader M200 (Tecan, Austria) at room temperature. The intensity of fluorescence per-gram of muscle tissue was expressed as arbitrary unit (a.u.). Extraction of G-Actin The content of G-actin was determined as described originally by Cano et al. (1991). Briefly, 1.0 g of muscle samples were homogenized with 8 mL of lysis buffer by a DY89-I high speed homogenizer (Scientz co., Ningbo, China) for 3 × 10 s at 10, 000 rpm while cooled in ice. The homogenate was centrifuged at 6, 000 g for 20 min at 4°C with a refrigerated centrifuge (Hunan Xiangyi Laboratory Instrument Development Co., Changsha, China). Suspensions were diluted with lysis buffer to the same protein concentration (mg/mL) by using a bicinchoninic acid (BCA) protein assay kit (Thermo Scientific, MA). The final supernatants were used for G-actin assays. Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) and Western Blotting The SDS-PAGE and western blotting were done to determine the levels of G-actin as described previously by Warner et al. (1997). Glyceraldehyde-3-phosphate dehydrogenas (GAPDH), with the molecular weight of 36 kDa, was used as standard reference for Western blot. G-actin sample (20 μg) was run on a 12.5% SDS-PAGE and then transferred to Polyvinylidene fluoride membranes (Immobilon-P, Millipore, MA). Membranes were blocked in 1 × PBST containing 5% fat-free dry milk for 1 h at 25°C and incubated with primary antibody for 24 h at 4°C. The primary antibodies of rabbit anti-beta-actin (ab8227, 1:5000, Abcam, Cambridge, MA) and mouse anti-GAPDH (ab8245, 1:5000, Abcam, Cambridge, MA) were diluted in 1 × PBST solution containing 0.5% fat-free dry milk, respectively. The goat anti-rabbit HRP-conjugated secondary antibodies (ab97051, 1:10,000, Abcam, Cambridge, MA) and goat anti-mouse HRP-conjugated secondary antibodies (ab97023, 1:10,000, Abcam, Cambridge, MA) were diluted in 1 × PBST solution containing 0.5% fat-free dry milk, respectively. According to instructions, the validation of the antibodies were suitable for western blot. Membranes were incubated with secondary antibodies at 25°C for 1 h. ECL Plus Western blotting detection kit (Amersham Biosciences, UK) was used to detect the blot according to the manufacturer's instructions. Western blot data were quantified by specialized software and a microcomputer (Quantity One Software, Bio-Rad Laboratories, PA). Statistical Analysis All results were expressed as means of three replications. All data were presented as mean ± standard error. Statistical analysis was carried out by the multiple-factors (treatments and storage time, including two-way interactions) analysis of variance (ANOVA) procedure (Duncan's Multiple Range Test) of SAS 8.0 software (SAS Institute Inc., Cary, NC) to analyze the change and the difference of shear force, cooking loss, MFI, particle size, F-actin contents and expression levels of G-actin among three treatments (control, 300 W ultrasound and 600 W ultrasound) during 168 h of storage. The significant level was set as 0.05. RESULTS AND DISCUSSION The Changes of Shear Force in three Groups The changes of shear force are shown in Figure 1A in three groups. The values of shear force significantly decreased during 24–168 h in control group and decreased during 8–168 h in 300 W and 600 W group (P < 0.05). Among three groups, the values of shear force in 600 W group were significantly lower during 24–168 h than 300 W group and control group (P < 0.05). The values of shear force in 300 W group were significantly lower at 24, 96 and 168 h than control group (P < 0.05). Our research indicated that UT reduced the shear force of goose meat. Figure 1. View largeDownload slide The changes of shear force (A) and cooking loss (B) in three groups during conditioning. a-i Identical letters indicated that there were no significant difference in different treatments (P > 0.05). Figure 1. View largeDownload slide The changes of shear force (A) and cooking loss (B) in three groups during conditioning. a-i Identical letters indicated that there were no significant difference in different treatments (P > 0.05). Consumer tenderness acceptability increased as shear force values decreased (Miller et al., 2001). Smith et al. (2002) observed that shear force values of bovine Semitendinosus muscle reduced due to ultrasonic treatment. It was reported that the cavitation generated by UT causes the physical disruption of meat (Alarconrojo et al., 2015). UT has been found to improve tenderness by the disruption of myofibrillar integrity (Got et al., 1999). The Change of Cooking Loss in three Groups The changes of cooking loss are shown in Figure 1B in three groups. The values of cooking loss significantly decreased during 24–168 h in control group, decreased during 8–168 h in 300 W group and decreased during 8–168 h in 600 W group (P < 0.05). Among three groups, the values of cooking loss in 600 W group were significantly lower during 24–168 h than 300 W group and control group (P < 0.05). The values of cooking loss in 300 W group were lower during 48–168 h than control group (P < 0.05). Our study indicated that UT improved the water-holding capacity of goose meat. Dolatowski et al. (2000) reported that UT changed the structure of meat tissue (especially protein molecules and cell) and increased the water-holding capacity of meat. Vimini et al. (2010) stated that UT could reduce cooking loss of meat, which was consistent with our results. It has been found that UT increased the water binding strength of meat which is responsible for improving cooking loss (Reynolds et al., 2010). Alarconrojo et al. (2015) assumed that ultrasound reduced cooking losses of meat by weakening connective tissues and myofibrillar structure. The Changes of MFI and Particle size in Three Groups The changes of MFI are shown in Figure 2 in three groups. The values of MFI significantly increased during 24–168 h in 300 W group and control group, during 8–168 h in 600 W group (P < 0.05). Among three treatments, samples in 600 W group showed the higher values of MFI at 48, 96 and 168 h than in 300 W group and control group (P < 0.05). The MFI values of 300 W group were significantly higher at 48, 96 and 168 h than control group (P < 0.05). Figure 2. View largeDownload slide The changes of myofibrillar fragmentation index in three groups during conditioning. a-j Identical letters indicated that there were no significant difference in different treatments (P > 0.05). Figure 2. View largeDownload slide The changes of myofibrillar fragmentation index in three groups during conditioning. a-j Identical letters indicated that there were no significant difference in different treatments (P > 0.05). Changes in D4,3, D3,2, Dv,0.1, Dv,0.5, Dv,0.9 of particle size in three groups during conditioning are shown in Table 1. The values of D4,3, D3,2, Dv,0.1 and Dv,0.9 significantly decreased during 24–168 h in the control while the value of Dv,0.5 in the control significantly decreased during 8–168 h (P < 0.05). The values of D4,3, D3,2, Dv,0.1 and Dv,0.5 significantly decreased during 24–168 h in the 300 W group, while the value of Dv,0.9 in the 300 W group significantly decreased during 8–168 h (P < 0.05). The values of D4,3, Dv,0.1, Dv,0.5 and Dv,0.9 significantly decreased during 8–168 h in the 600 W group, while the value of D3,2 in the 600 W group significantly decreased during 24–168 h (P < 0.05). Among three groups, the values of D4,3, D3,2, Dv,0.1 and Dv,0.5 in 600 W group were lower during 48–168 h than 300 W group and control group, while the value of Dv,0.9 in 600 W group were significantly lower during 24–168 h than 300 W group and control group (P < 0.05). The values of D3,2, Dv,0.1 and Dv,0.5 in 300 W group were significantly lower during 48–168 h than the control and the value of D4,3 in 300 W group were significantly lower during 48–96 h than the control, while the value of Dv,0.9 in 300 W group were significantly lower during 96–168 h than control group (P < 0.05). The particle size distributions showed that UT increased the level of smaller particles and decreased the level of large particles compared to the control. Table 1. Changes in average particle size of myofibrillar proteins in three groups during conditioning.   D4,3/μm  D3,2/μm  DV,0.1/μm  DV,0.5/μm  DV,0.9/μm  CK-0h  196.55 ± 1.48a  61.7 ± 1.33a  32.83 ± 1.12a  140.36 ± 1.14a  423.25 ± 2.98a  CK-8h  195.27 ± 1.17a,b  59.38 ± 1.23a,b  31.84 ± 1.26a,b  138.31 ± 0.76a  418.91 ± 2.39a,b  CK-24h  193.41 ± 1.28b  57.27 ± 1.33b,c  30.45 ± 1.28b  135.68 ± 1.32b  412.53 ± 2.34b,c  CK-48h  188.01 ± 1.2d  52.2 ± 1.75d  28.67 ± 0.96c  129.46 ± 0.88c  398.57 ± 3.99d,e  CK-96h  183.45 ± 1.11f  47.43 ± 1.38e,f  25.42 ± 1.15d  124.02 ± 1.17d,e  385.38 ± 3.34e,f  CK-168h  179.33 ± 1.02 g,h  43.39 ± 1.35g  22.6 ± 1.14f  116.5 ± 0.89f  372.34 ± 2.88h  300W-0h  196 ± 0.54a  60.71 ± 1.4a,b  32.42 ± 0.6a,b  139.22 ± 0.91a  421.5 ± 2.38a,b  300W-8h  194.89 ± 0.37a,b  58.42 ± 1.18b  31.21 ± 0.65a,b  137.38 ± 0.73a,b  414.71 ± 2.26b  300W-24h  192.67 ± 0.8b,c  56.45 ± 1.35b,c  29.63 ± 1.27b,c  132.4 ± 1.11b,c  407.48 ± 2.59c  300W-48h  185.78 ± 1.09e  49.46 ± 1.26e  26.24 ± 0.79d  126.69 ± 0.83d  391 ± 2.46e  300W-96h  180.79 ± 1.2g  43.22 ± 1.12g  23.93 ± 0.91e  116.5 ± 1.09f  378.64 ± 2.49g  300W-168h  176.5 ± 1.38h  40.14 ± 1.14h  20.6 ± 0.98g  110.72 ± 1.15g  365.63 ± 1.2i  600W-0h  195.3 ± 0.82a,b  60.45 ± 1.23a,b  31.93 ± 0.85a,b  138.45 ± 0.77a  418.91 ± 1.09a,b  600W-8h  193.7 ± 0.79b  58.14 ± 1.09b,c  30.4 ± 1.07b  136.54 ± 1.31a,b  412.67 ± 1.64b,c  600W-24h  190.7 ± 0.72c  55.47 ± 1.5c  28.33 ± 1.04c  130.21 ± 1.24c  400.96 ± 1.75d  600W-48h  181.67 ± 0.88g  47.43 ± 1.06f  24.73 ± 1e  123.22 ± 0.89e  383.64 ± 2.06f  600W-96h  177.19 ± 0.83h  41.25 ± 1.11h  22.42 ± 0.96f  111.59 ± 1.48g  371.45 ± 1.54h  600W-168h  174.45 ± 0.92i  38.15 ± 1.34i  19.43 ± 0.9h  105.63 ± 1.58h  359.57 ± 2.45j    D4,3/μm  D3,2/μm  DV,0.1/μm  DV,0.5/μm  DV,0.9/μm  CK-0h  196.55 ± 1.48a  61.7 ± 1.33a  32.83 ± 1.12a  140.36 ± 1.14a  423.25 ± 2.98a  CK-8h  195.27 ± 1.17a,b  59.38 ± 1.23a,b  31.84 ± 1.26a,b  138.31 ± 0.76a  418.91 ± 2.39a,b  CK-24h  193.41 ± 1.28b  57.27 ± 1.33b,c  30.45 ± 1.28b  135.68 ± 1.32b  412.53 ± 2.34b,c  CK-48h  188.01 ± 1.2d  52.2 ± 1.75d  28.67 ± 0.96c  129.46 ± 0.88c  398.57 ± 3.99d,e  CK-96h  183.45 ± 1.11f  47.43 ± 1.38e,f  25.42 ± 1.15d  124.02 ± 1.17d,e  385.38 ± 3.34e,f  CK-168h  179.33 ± 1.02 g,h  43.39 ± 1.35g  22.6 ± 1.14f  116.5 ± 0.89f  372.34 ± 2.88h  300W-0h  196 ± 0.54a  60.71 ± 1.4a,b  32.42 ± 0.6a,b  139.22 ± 0.91a  421.5 ± 2.38a,b  300W-8h  194.89 ± 0.37a,b  58.42 ± 1.18b  31.21 ± 0.65a,b  137.38 ± 0.73a,b  414.71 ± 2.26b  300W-24h  192.67 ± 0.8b,c  56.45 ± 1.35b,c  29.63 ± 1.27b,c  132.4 ± 1.11b,c  407.48 ± 2.59c  300W-48h  185.78 ± 1.09e  49.46 ± 1.26e  26.24 ± 0.79d  126.69 ± 0.83d  391 ± 2.46e  300W-96h  180.79 ± 1.2g  43.22 ± 1.12g  23.93 ± 0.91e  116.5 ± 1.09f  378.64 ± 2.49g  300W-168h  176.5 ± 1.38h  40.14 ± 1.14h  20.6 ± 0.98g  110.72 ± 1.15g  365.63 ± 1.2i  600W-0h  195.3 ± 0.82a,b  60.45 ± 1.23a,b  31.93 ± 0.85a,b  138.45 ± 0.77a  418.91 ± 1.09a,b  600W-8h  193.7 ± 0.79b  58.14 ± 1.09b,c  30.4 ± 1.07b  136.54 ± 1.31a,b  412.67 ± 1.64b,c  600W-24h  190.7 ± 0.72c  55.47 ± 1.5c  28.33 ± 1.04c  130.21 ± 1.24c  400.96 ± 1.75d  600W-48h  181.67 ± 0.88g  47.43 ± 1.06f  24.73 ± 1e  123.22 ± 0.89e  383.64 ± 2.06f  600W-96h  177.19 ± 0.83h  41.25 ± 1.11h  22.42 ± 0.96f  111.59 ± 1.48g  371.45 ± 1.54h  600W-168h  174.45 ± 0.92i  38.15 ± 1.34i  19.43 ± 0.9h  105.63 ± 1.58h  359.57 ± 2.45j  a-jIdentical letters indicate that there is no significant difference in different processing points (P > 0.05). View Large Table 1. Changes in average particle size of myofibrillar proteins in three groups during conditioning.   D4,3/μm  D3,2/μm  DV,0.1/μm  DV,0.5/μm  DV,0.9/μm  CK-0h  196.55 ± 1.48a  61.7 ± 1.33a  32.83 ± 1.12a  140.36 ± 1.14a  423.25 ± 2.98a  CK-8h  195.27 ± 1.17a,b  59.38 ± 1.23a,b  31.84 ± 1.26a,b  138.31 ± 0.76a  418.91 ± 2.39a,b  CK-24h  193.41 ± 1.28b  57.27 ± 1.33b,c  30.45 ± 1.28b  135.68 ± 1.32b  412.53 ± 2.34b,c  CK-48h  188.01 ± 1.2d  52.2 ± 1.75d  28.67 ± 0.96c  129.46 ± 0.88c  398.57 ± 3.99d,e  CK-96h  183.45 ± 1.11f  47.43 ± 1.38e,f  25.42 ± 1.15d  124.02 ± 1.17d,e  385.38 ± 3.34e,f  CK-168h  179.33 ± 1.02 g,h  43.39 ± 1.35g  22.6 ± 1.14f  116.5 ± 0.89f  372.34 ± 2.88h  300W-0h  196 ± 0.54a  60.71 ± 1.4a,b  32.42 ± 0.6a,b  139.22 ± 0.91a  421.5 ± 2.38a,b  300W-8h  194.89 ± 0.37a,b  58.42 ± 1.18b  31.21 ± 0.65a,b  137.38 ± 0.73a,b  414.71 ± 2.26b  300W-24h  192.67 ± 0.8b,c  56.45 ± 1.35b,c  29.63 ± 1.27b,c  132.4 ± 1.11b,c  407.48 ± 2.59c  300W-48h  185.78 ± 1.09e  49.46 ± 1.26e  26.24 ± 0.79d  126.69 ± 0.83d  391 ± 2.46e  300W-96h  180.79 ± 1.2g  43.22 ± 1.12g  23.93 ± 0.91e  116.5 ± 1.09f  378.64 ± 2.49g  300W-168h  176.5 ± 1.38h  40.14 ± 1.14h  20.6 ± 0.98g  110.72 ± 1.15g  365.63 ± 1.2i  600W-0h  195.3 ± 0.82a,b  60.45 ± 1.23a,b  31.93 ± 0.85a,b  138.45 ± 0.77a  418.91 ± 1.09a,b  600W-8h  193.7 ± 0.79b  58.14 ± 1.09b,c  30.4 ± 1.07b  136.54 ± 1.31a,b  412.67 ± 1.64b,c  600W-24h  190.7 ± 0.72c  55.47 ± 1.5c  28.33 ± 1.04c  130.21 ± 1.24c  400.96 ± 1.75d  600W-48h  181.67 ± 0.88g  47.43 ± 1.06f  24.73 ± 1e  123.22 ± 0.89e  383.64 ± 2.06f  600W-96h  177.19 ± 0.83h  41.25 ± 1.11h  22.42 ± 0.96f  111.59 ± 1.48g  371.45 ± 1.54h  600W-168h  174.45 ± 0.92i  38.15 ± 1.34i  19.43 ± 0.9h  105.63 ± 1.58h  359.57 ± 2.45j    D4,3/μm  D3,2/μm  DV,0.1/μm  DV,0.5/μm  DV,0.9/μm  CK-0h  196.55 ± 1.48a  61.7 ± 1.33a  32.83 ± 1.12a  140.36 ± 1.14a  423.25 ± 2.98a  CK-8h  195.27 ± 1.17a,b  59.38 ± 1.23a,b  31.84 ± 1.26a,b  138.31 ± 0.76a  418.91 ± 2.39a,b  CK-24h  193.41 ± 1.28b  57.27 ± 1.33b,c  30.45 ± 1.28b  135.68 ± 1.32b  412.53 ± 2.34b,c  CK-48h  188.01 ± 1.2d  52.2 ± 1.75d  28.67 ± 0.96c  129.46 ± 0.88c  398.57 ± 3.99d,e  CK-96h  183.45 ± 1.11f  47.43 ± 1.38e,f  25.42 ± 1.15d  124.02 ± 1.17d,e  385.38 ± 3.34e,f  CK-168h  179.33 ± 1.02 g,h  43.39 ± 1.35g  22.6 ± 1.14f  116.5 ± 0.89f  372.34 ± 2.88h  300W-0h  196 ± 0.54a  60.71 ± 1.4a,b  32.42 ± 0.6a,b  139.22 ± 0.91a  421.5 ± 2.38a,b  300W-8h  194.89 ± 0.37a,b  58.42 ± 1.18b  31.21 ± 0.65a,b  137.38 ± 0.73a,b  414.71 ± 2.26b  300W-24h  192.67 ± 0.8b,c  56.45 ± 1.35b,c  29.63 ± 1.27b,c  132.4 ± 1.11b,c  407.48 ± 2.59c  300W-48h  185.78 ± 1.09e  49.46 ± 1.26e  26.24 ± 0.79d  126.69 ± 0.83d  391 ± 2.46e  300W-96h  180.79 ± 1.2g  43.22 ± 1.12g  23.93 ± 0.91e  116.5 ± 1.09f  378.64 ± 2.49g  300W-168h  176.5 ± 1.38h  40.14 ± 1.14h  20.6 ± 0.98g  110.72 ± 1.15g  365.63 ± 1.2i  600W-0h  195.3 ± 0.82a,b  60.45 ± 1.23a,b  31.93 ± 0.85a,b  138.45 ± 0.77a  418.91 ± 1.09a,b  600W-8h  193.7 ± 0.79b  58.14 ± 1.09b,c  30.4 ± 1.07b  136.54 ± 1.31a,b  412.67 ± 1.64b,c  600W-24h  190.7 ± 0.72c  55.47 ± 1.5c  28.33 ± 1.04c  130.21 ± 1.24c  400.96 ± 1.75d  600W-48h  181.67 ± 0.88g  47.43 ± 1.06f  24.73 ± 1e  123.22 ± 0.89e  383.64 ± 2.06f  600W-96h  177.19 ± 0.83h  41.25 ± 1.11h  22.42 ± 0.96f  111.59 ± 1.48g  371.45 ± 1.54h  600W-168h  174.45 ± 0.92i  38.15 ± 1.34i  19.43 ± 0.9h  105.63 ± 1.58h  359.57 ± 2.45j  a-jIdentical letters indicate that there is no significant difference in different processing points (P > 0.05). View Large MFI is one of the most widely useful indicator to determine postmortem proteolysis and tenderization in meat (Volpelli et al., 2005). Kang et al. (2017) reported that beef treated with ultrasound could increase MFI values thus increase tenderness of meat. Karumendu et al. (2009) demonstrated that the increase in MFI is accompanied by a decrease in shear force. In our study, the increase of MFI by UT contributed to the decrease of Warner-Bratzler shear force values and cooking loss during conditioning. Shin et al. (2008) indicated that the decrease of particle size of myofibril proteins contributed to meat tenderization. Li et al. (2014) reported that UT (450 W) caused a decrease in particle size of myofibril fragmentation in chicken breast meat since the distribution became more uniform and narrowed. The reduction of particle size is due to proteins dissociation during cavitation by UT and particles under sonication were violently agitated which resulted in broken aggregate particles (Zhang et al., 2017). It demonstrated that the reduction of particle size for myofibril fragmentation was contributed to the decrease of shear force values at storage periods directly (Crouse et al., 1991). The smaller particle size contributed to stronger protein-water interactions which strengthened proteins solubility and water holding capacity (Jambrak et al., 2014). The decrease in particle size by UT was due to the disruption of myofibril structure (Jayasooriya et al., 2004). In our research, the decrease in particle size by UT was related to the result of the increase in MFI. The Morphological Change of Actin Filaments in Three Groups The changes of actin filaments by FITC-phalloidin staining in three groups are shown in Figure 3. Actin filaments morphology did not show significant fracture among three treatments at 0 h. In the control, actin filaments disrupted slightly during 96–168 h. Actin filaments structure in 300 W group changed more dramatically during 96–168 h than the control. Actin filaments in 600 W group disrupted more dramatically during 96–168 h than 300 W group and the control. It indicated that ultrasound treatment accelerated the proteolysis of actin filaments. Figure 3. View largeDownload slide The changes of actin filaments in three groups during conditioning. CK, 300 W and 600 W indicated control, 300 W ultrasound group and 600 W ultrasound group, respectively. Images were obtained using a fluorescence microscope with the 100 × objective (Olympus, Japan). Arrows in 300 W and 600 W groups implied that the actin filaments were disrupted dramatically. Figure 3. View largeDownload slide The changes of actin filaments in three groups during conditioning. CK, 300 W and 600 W indicated control, 300 W ultrasound group and 600 W ultrasound group, respectively. Images were obtained using a fluorescence microscope with the 100 × objective (Olympus, Japan). Arrows in 300 W and 600 W groups implied that the actin filaments were disrupted dramatically. FITC-phalloidin enabled us to observe the distribution of F-actin in myofibril structure (Schmit and Lambert, 1990). The basic contractile unit of myofibrils is the sarcomere which include myosin, actin and nebulin filament (Littlefield and Fowler, 2008). Meat treated with ultrasound resulted in the disintegration of the myofibrillar scaffold (Chen et al., 2015). Ahn et al. (2003) thought that the increase of MFI with aging may be related to the breaking of myofibrillar proteins into segments near the Z-disk. Latoch (2010) indicated that UT can induce the degradation of I-Z-I band proteins and then increase the MFI. It was noted that actin accounts for 22% of myofibrillar protein and that its content was higher than the level of some cytoskeletal proteins in Z-disk substantially (Yates and Greaser, 1983). We thought that the degradation of cytoskeletal proteins near the Z-disk was not the directly reason for the increase of MFI. Our result indicated that the disruption of actin filaments but not Z-disks by UT could directly lead to the increase of MFI and the decrease of particle size. It could be an important factor in meat tenderization during conditioning. Changes of F-Actin and G-Actin Contents in Three Treatments The changes of F-actin content in myofibrils are shown in Figure 4. The content of F-actin in all treatments had the highest values at 0 h which indicated a very low extent of F-actin depolymerization. The content of F-actin significantly reduced during 8–168 h in the control and 300 W group and decreased during 0–168 h in 300 W group (P < 0.05). 600 W group showed lower content of F-actin during 8–168 h compared to the control and 300 W group (P < 0.05). Compared to control group, the content of F-actin was significantly lower during 24–168 h in 300 W group (P < 0.05). Figure 4. View largeDownload slide The changes of F-actin content in myofibrils in three groups during conditioning. a-j Identical letters indicated that there were no significant difference in different treatments (P > 0.05). Figure 4. View largeDownload slide The changes of F-actin content in myofibrils in three groups during conditioning. a-j Identical letters indicated that there were no significant difference in different treatments (P > 0.05). The changes of G-actin (43 kDa) levels are shown in Figure 5 by western blot analysis in three groups. The G-actin bands significantly increased during 0–24 h and during 96–168 h in control group, during 24–96 h in 300 W group and 600 W group (P < 0.05), respectively. The G-actin bands in 600 W group had larger values from 24 to 96 h than in the control and 300 W group (P < 0.01). The G-actin bands of 300 W group had higher values during 96–168 h than the control (P < 0.05). The decrease of F-actin and increase of G-actin implied that F-actin depolymerized to G-actin. The rate of the depolymerization of F-actin to G-actin in 600 W group was faster than 300 W group; the rate in the control was the slowest among three groups. Figure 5. View largeDownload slide The changes of G-actin levels by SDS-PAGE and western blot in three groups during conditioning. a-e Identical letters in the line indicated that there were no significant difference in different treatments (P > 0.05). Figure 5. View largeDownload slide The changes of G-actin levels by SDS-PAGE and western blot in three groups during conditioning. a-e Identical letters in the line indicated that there were no significant difference in different treatments (P > 0.05). The phallacidin-stained F-actin of tissues and cells is widely used to quantify the content of F-actin (Sampath and Pollard, 1991). UT lead to a decrease in the expression of cytoskeletal proteins such as F-actin by immunocytochemical staining (Monici et al., 2007). Saleem et al. (2015) reported that ultrasonication causes the structural changes in myosin and actin of chicken breast muscle. It has been shown that nebulin participates in the stabilization of F-actin (Wang and Wright, 1988). Xiong et al. (2012) demonstrated that nebulin were degraded from chicken breast muscle by UT. It indicated that ultrasound could weaken F-actin by degraded nebulin proteins. Littlefield et al. (2001) indicated that there is an actin dynamics fluctuating between actin filaments and monomers in skeletal muscle cells. It is known that treatment with ultrasound (sonication) reinforce actin depolymerization (Carlier et al., 1984). UT could release Ca2+ which may regulated the pathway of the actin depolymerize to G-actin (Lange and Brandt, 1996). G-actin can be prepared from F-actin by the treatment of ultrasonic (Mihashi, 1964). A slow actin depolymerization began to occur after sonication, indicating that its alteration is caused by ultrasound extraction (Wendel and Dancker, 1986). In our results, we found that ultrasound treatments accelerated the transformation from F-actin to G-actin. CONCLUSIONS The disruption of actin filaments but not Z-disks by UT could contribute to myofibrillar fraction directly and the tenderization of goose meat. UT accelerated the transformation from F-actin to G-actin. 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Poultry ScienceOxford University Press

Published: Apr 20, 2018

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