The preservable effects of ultrasound-assisted alginate oligosaccharide soaking on cooked crayfish subjected to Freeze-Thaw cycles

Graphical abstract

Highlights

• •Freeze-thaw cycles reduced water retention of cooked crayfish.
• •Ultrasound-assisted alginate oligosaccharide soaking improved water status.
• •It also mitigated the damage of tissue and reduced ice crystal size.

Abstract

To improve the quality of cooked and frozen crayfish after repeated freeze–thaw cycles, the effects of alginate oligosaccharide (1 %, w/v) with ultrasound-assisted (40 W, 3 min) soaking (AUS) on the physicochemical properties were investigated. The AUS samples improved water-holding capacity with 19.47 % higher than the untreated samples. Low-field nuclear magnetic resonance confirmed that mobile water (T₂₂) in the samples after 5 times of freeze–thaw cycles was reduced by 13.02 % and 29.34 % with AUS and without treatment, correspondingly; and with AUS and without treatment, average size of the ice crystals was around 90.26 μm² and 113.73 μm², and average diameter of the ice crystals was 5.83 μm and 8.14 μm, respectively; furthermore, it enhanced the solubility and zeta potential, lowered the surface hydrophobicity, reduced the particle size, and maintained the secondary and tertiary structures of myofibrillar protein (MP) after repeated freeze-thawing. Gel electrophoresis revealed that the AUS treatment mitigated the denaturation of MPs. Scanning electron microscopy revealed that the AUS treatment preserved the structure of the tissue. These findings demonstrated that the AUS treatment could enhance the water retention and physicochemical properties of protein within aquatic meat products during temperature fluctuations..

1. Introduction

Crayfish (Procambarus clarkii), with high protein content (16–20 %) and low-fat content (lower than 2 %), exhibits rapid growth, and is widely farmed in the central regions of China. It has become one of the major farmed economic species in the freshwater aquaculture with annual output of around 2.64 million tons [1], [2]. As a seasonal product, fresh crayfish serves as for instant meals since it contains a large amount of moisture (nearly accounted for 75 % of total muscle meat) and microorganisms (including Salmonella, Staphylococcus aureus, and Vibrio parahaemolyticus), which promotes enzymatic and oxidative reactions, and eventually destroys the value of crayfish [3], so excessive crayfish (approximately-one third of total products) needs to be processed and stored for year-round consumption, which is usually cooked and transported in the frozen state [4]. However, imperfect cold-chain logistics with temperature fluctuation makes frozen meat repeatedly thaw and freeze during transportation, which seriously affects the nourishment and safety of meat [5]. It has been reported that as the number of freeze–thaw cycles increases, the drip loss of fish balls occurs significantly, exacerbating the mechanical damage of the fish balls [6].

Water-retention agents, such as sugar, starch, alginate, konjac glucomannan, and carrageenan, have been widely applied as to reduce drip loss in food products during storage [7]. While these biomacromolecules can be added during the processing of restructured food, such as surimi gel products, it is difficult to penetrate natural muscle tissue, such as shrimp and crayfish. For this reason, oligosaccharides with smaller molecular weight and lower steric hindrance are being gradually employed in frozen food storage. It has been reported that xylooligosaccharide could provide better cryoprotective function to porcine myofibrillar proteins (MPs) than other sugars [8]. Zhang et al. [9] impregnated frozen peeled shrimp (Litopenaeus vannamei) with carrageenan oligosaccharide, significantly improving the water-holding capacity (WHC). This was ascribed to the binding of the oligosaccharide with proteins to capture free water in muscle tissues during frozen storage. However, although numerous studies have investigated the effect of cryoprotective agents on the quality of frozen food products, there is limited data available pertaining to the mechanism behind the cryoprotective activities of oligosaccharide during long-term frozen storage with temperature fluctuations. Moreover, with the advantages of being non-invasive, directional, non-polluting, and penetrating, ultrasound has also become a popular processing technology to improve the flavor, tenderness, and other aspects of food quality [10], [11], [12]. Ultrasound waves propagate through a liquid medium as a series of compression waves, generating alternating regions of high and low pressure, and the rapid transition between compression and dilation results in the generation of cavitation bubbles [13]. Therefore, ultrasonic-assisted soaking may accelerate the diffusion of a water-retention agent into the interior of crayfish so that the agent could adequately mix with the tissues.

Alginate oligosaccharide, a biopolymer widely used in the food, pharmaceutical, and chemical industries, possesses good solubility and bioavailability, and exhibits a variety of biological activities, including antioxidant, immunomodulatory, antibacterial, and anti-inflammatory activities. However, unlike other oligosaccharides, research on the effect of alginate oligosaccharide on the quality of frozen aquatic products has been limited. In this study, ultrasound-assisted alginate oligosaccharide soaking was applied to cooked crayfish to investigate its influence on ice crystal growth and microstructural changes, as well as water retention after freeze–thaw cycles, which may provide a novel method to stabilize the quality of crayfish products during the temperature fluctuation.

2. Materials and methods

2.1. Materials

Fresh crayfish (P. clarkia, 25 ± 2 g) was purchased from a local market (Nanjing, China). Sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) loading buffer, precast SDS-PAGE gel (12 %, 10-well), and Real Band 3-color regular range (5–245 kDa) protein marker, were purchased from Sangon Biotech Co., Ltd. (Shanghai, China). Alginate oligosaccharide (Mw ≈ 2800 Da) was provided by Qingdao Oligo Biotech Co., Ltd (Qingdao, China). Other reagents and chemicals of analytical grade were purchased from Sinopharm Group Co., Ltd. (Shanghai, China).

2.2. Preparation of crayfish samples

Fresh crayfish (25 ± 2 g) was rinsed with water to remove surface deposits and divided into four groups. Samples of the first group were shelled and is denoted as CK. The second group, denoted as CED, was cooked for 15 min at 100 °C with intermittent mild stirring and then shelled. CED soaked in a 1 % alginate oligosaccharide solution (the concentration was selected by pre-experiments) for 1 h is denoted as AOS. CED soaked for 1 h in 1 % alginate oligosaccharide assisted with ultrasound (40 W, 3 min, optimized conditions got by the result in Table S1) is named AUS. All samples except CK underwent 0, 1, 2, and 5freeze–thaw cycles, marked as 0F-T, 1F-T, 2F-T, and 5F-T, respectively. A freeze–thaw cycle entailed freezing at −20 °C for 12 h and thawing at 4 °C for 3 h.

2.3. Measurement of WHC

Approximately 2 g of a crayfish sample was wrapped in absorbent cotton and centrifuged for 10 min (4 °C, 9000 r/min). The WHC of crayfish was calculated using the following equation:

$$\mathit{WHC}(\%)=1-\frac{m_1-m_2}{m_1}\times 100$$ (1)

where m₁ and m₂ are the weights of crayfish before and after centrifugation, respectively.

2.4. Low-field nuclear magnetic resonance and magnetic resonance imaging

Low-field nuclear magnetic resonance (LF NMR) and magnetic resonance imaging (MRI) analyses of water status in crayfish were conducted using a MesoMR23-060H-I NMR imaging and analysis system (Suzhou Niumag Analytical Instrument Co., Ltd., Suzhou, China). The crayfish was placed in a glass tube (outer diameter: 60 mm) and inserted into the NMR probe. The spin–spin relaxation time of water was measured based on Carr-Purcell-Meiboom-Gill pulse sequences. The test settings were as follows: a 14 μs 90° pulse followed by a 26.48 μs 180° pulse; waiting time, 3500 ms; repeated scan number, 16; echo count, 15,000; and echo time, 200 μs. The hydrogen proton density and distribution in crayfish were determined using the following MRI parameters: echo time, 20 ms; repetition time, 500 ms; read size × phase size, 256 × 192; average number, 4; and field of view (FOV), 100 × 100 mm².

2.5. Measurement of ice crystal size

The size of ice crystals in cooked crayfish after repeated freeze–thaw cycles was determined following the method described by Liu et al. [14] with some modifications. Briefly, samples were cut into small pieces and soaked overnight in 4 % paraformaldehyde (1:10, w/v). After embedding, the samples were subjected to frozen sectioning and sliced into 8 μm-thick sections along the muscle fibers. The sections were sequentially placed in xylene for 20 min, anhydrous ethanol for 10 min, 75 % ethanol for 5 min, and washed with distilled water. The sections were then placed in hematoxylin staining solution for 5 min, followed by fractionation solution and distilled water. The sections were then dehydrated in 85 % and 95 % alcohol for 5 min each and stained with eosin staining solution for 5 min. Finally, the sections were placed in anhydrous ethanol for 15 min and then in xylene for 10 min and sealed with neutral gum. Microscopy was carried out with an optical microscope (Nikon Eclipse E100, Japan) and the ice crystal size of cooked crayfish after repeated freeze–thaw cycles was calculated using Image Pro Plus 6.0 software (Media Cybernetics, Silver Spring, MD, USA).

2.6. Solubility, particle size, zeta potential, and surface hydrophobicity of MPs extracted from crayfish

MPs were extracted from cooked crayfish samples according to the method described by Shi et al. [15] and dispersed in phosphate buffer (0.6 M KCl, 10 mM K₂HPO₄, pH 6.0) to determine the protein concentration via the biuret method. The solubility of the MPs was determined according to Zhi et al. [16]. Briefly, the MPs solution was diluted to 2.5 mg/mL with phosphate buffer (20 mM, pH 6.0) and then centrifuged for 10 min (10,000 g, 4 °C). The supernatant was collected, and the ratio of the supernatant protein concentration to 2.5 mg/mL was the MP solubility.

The particle size and zeta potential of the MPs were determined according to the method described by Gao et al. [17] with minor modifications as following: The experiments were performed at room temperature and the MPs solution was adjusted to 1 mg/mL with phosphate buffer (20 mM, pH 6.0). 1 mL of the diluted MPs sample was placed in a Malvern cuvette, and the particle size and zeta potential of MPs were determined using a nanoparticle size and zeta potential analyzer (Nicomp-Z3000, PSS, USA).

The surface hydrophobicity of MPs was determined using bromophenol blue as a hydrophobic probe according to the method described by Hu et al. [18]. The MPs solution was diluted to 2.5 mg/mL with phosphate buffer (20 mM, pH 6.0) and mixed with 0.2 mL of a bromophenol blue solution (1 mg/mL), with a solution containing 1 mL of phosphate buffer and 0.2 mL of the bromophenol blue (1 mg/mL) as control. All samples were incubated at 25 °C for 10 min with shaking and then centrifuged for 15 min (2000 g, 4 °C). Finally, 0.4 mL of the supernatant was mixed with 3.6 mL phosphate buffer (20 mM, pH 6.0), and the absorbance (A_sample) was measured at 595 nm (UV-2102C spectrophotometer, Unico, USA). These steps were repeated with phosphate solution without MPs and the control absorbance (A_control) was measured. The surface hydrophobicity of the MPs was calculated as follows:

$$\mathit{Surface}hydrophobicity\left(\mu g\right)=\frac{200\times \left(A_{\mathit{Control}}-A_{\mathit{Sample}}\right)}{A_{\mathit{Control}}}$$ (2)

2.7. Raman spectroscopy

Laser Raman spectroscopy of the cooked crayfish samples was performed using a Raman spectrometer (DXR532; Thermo Fisher Scientific, USA) with a 785 nm argon laser. Data was collected in the scanning range of 400–4000 cm⁻¹ with a spectral resolution of 3 cm⁻¹. Peakfit software (version 4.12, SeaSolve Software, Inc., Framingham, MA, USA) was used to analyze the secondary structure of the proteins in the cooked crayfish samples and expressed as a percentage of α-helix, β-sheet, β-turn, and random coil conformations.

2.8. Endogenous fluorescence analysis of MPs extracted from crayfish

The intrinsic fluorescence intensities of the cooked crayfish samples were determined by fluorescence spectroscopy using a Cytation 3 Cell imaging multimode microplate reader (BioTek Instruments, Winooski, USA). The concentration of MPs was adjusted to 500 μg/mL and placed in an opaque 96-well plate. The excitation wavelength was 280 nm and the emission spectra measured in the range of 300–430 nm. The excitation and emission slit widths were 2.5 nm, and the scanning speed was 2 nm/s.

2.9. Sodium dodecyl sulfate–polyacrylamide gel electrophoresis

SDS-PAGE was performed to determine structural changes in the MPs [19]. Briefly, the MPs concentration was adjusted to 1 mg/mL and mixed with loading buffer at a ratio of 1:1 (v/v). The mixture was then heated at 95 °C for 10 min and centrifuged for 5 min (10,000 r/min). 20 μL of the supernatant and 10 μL of the marker were added to the loading well. Electrophoretic buffer (50 mM Tris, 50 mM HEPES, 0.1 % SDS, and 2 mM EDTA) was added, and electrophoresis was conducted at 150 V for 40 min. Afterwards, the gel was soaked for 4 h in Coomassie Brilliant Blue R-250 solution for staining, then destained in a solution containing 5 % ethanol and 10 % glacial acetic acid. An Image Lab gel imaging system (Tanon 2500; Tanon Science & Technology, Co., Ltd., Shanghai, China) was used to photograph the electrophoretic gels.

2.10. Scanning electron microscopy

The cooked crayfish samples were cut into small cubes and soaked overnight in a 2.5 % glutaraldehyde solution. The samples were then washed with 0.1 M phosphate buffer (pH 7.2) for 15 min four times. After being dehydrated with 50 %, 70 %, 80 %, 90 %, and 100 % ethanol solutions for 10 min each, the samples were freeze-dried and then sputter-coated with gold for observation by Scanning electron microscopy (SEM, EVO-LS10, Oberkochen, Germany).

2.11. Statistical analysis

The experiments were conducted in triplicate, with the results expressed as the mean and standard deviation. The data were analyzed by one-way analysis of variance, and the significant difference between the mean values was determined by Duncan’s multiple range test (p < 0.05) using SPSS statistical software (version 19.0, SPSS, Inc., Chicago, IL, USA).

3. Results and discussion

3.1. WHC and water status of crayfish meat

Fig. 1a shows that the WHC of all groups decreased significantly with increasing freeze–thaw cycles (p < 0.05) compared with CK. After five freeze–thaw cycles, the WHC of the CED, AOS, and AUS samples decreased by 12.72 %, 11.22 %, and 9.30 %, compared to their initial values, respectively. When cooked crayfish is stored at −20 °C, the water in crayfish samples froze, and, as the number of freeze–thaw cycles increased to 5 times, the initially small ice crystals developed into larger ones, average size of the ice crystals increased from around 42.35 μm² to 113.73 μm², and average diameter of the ice crystals expanded from approximately 5.82 μm to 8.14 μm, which can be seen in Fig. 2. This resulted in mechanical damage to the tissue structure, manifesting as increased muscle fiber bundle gaps, destruction of the structure, and a reduction in the WHC of the crayfish. However, the WHC of the AUS sample was 19.47 % and 15.37 % higher than those of CED and AOS, respectively, which initially confirmed that ultrasound-assisted soaking in alginate oligosaccharide could improve the WHC to some extent. This can be attributed to the formation of cavities by the cavitation effect, which enabled the penetration of water combining with small molecules such as oligosaccharide, thus increasing the WHC [20]. This was consistent with the observations of Li et al. [21], who studied the effects of ultrasound treatment on the quality of chicken breast meat and found that there were more cavities in the chicken breast after ultrasound treatment, which significantly improved the textural properties and WHC.

Fig. 1.

Changes in water holding capacity (a), transverse relaxation time (T₂, b) and proton density images (c) of cooked crayfish subjected to different treatments. Different letters in the same column of the same indicator showed significant difference (p < 0.05).

Fig. 2.

Changes in morphology (a, scale bar represents 50 μm), average diameter (b) and area (c) of ice crystal of cooked crayfish subjected to different treatments. Different letters in the same column of the same indicator showed significant difference (p < 0.05).

To gain insight into the changes in water distribution and dynamic water status of cooked crayfish under different treatments, the T₂ inversion spectrum was obtained, revealing three peaks corresponding to bound (T₂₁), immobilized (T₂₂), and free (T₂₃) water, shown in Fig. 1b. There was no significant change in the T₂₁ values with increasing freeze–thaw cycles (p > 0.05); however, there was a significant difference between the T₂₁ values of the different treatments (p < 0.05). After five freeze–thaw cycles, the T₂₂ peak of CED, AOS, and AUS right-shifted to 115.90 ms, 139.66 ms, and 188.60 ms, respectively, correspondingly the values decreased by 29.34 %, and 14.86 % and 13.02 % (Table S2). Li et al. [22] reported that ice crystals formed by repeated freezing and thawing caused physical damage to the structure of MPs, resulting in a larger external surface and a decrease in immobilized water content. Furthermore, ultrasound treatment promotes the rapid penetration of alginate oligosaccharide into the tissue, which can combine with more free water and convert it into immobilized water, leading to the increase of the WHC as was mentioned above.

Changes in moisture distribution in crayfish was observed using MRI. In the pseudo-color image, red areas indicate high moisture content (high density of hydrogen atoms), whereas blue areas indicate low moisture content (low density of hydrogen atoms). As shown in Fig. 1c, the moisture content of the outer layer is initially higher than that of the inner layer. With increasing freeze–thaw cycles, the color of the pseudo-color image fades, indicating that moisture migrated from the outer to the inner layer and, eventually, the overall moisture content decreased. The MRI results indicate that the moisture content of the AUS sample was higher than that of the other treatment groups, corroborating the LF NMR results. This was likely due to the high energy released by ultrasound, which created micro-jets on the surface, resulting in smaller ice crystals through accelerated heat transfer, thus preserving the original quality characteristics of the frozen food [23]. Sun et al. [3] found that the water holding capacity of crayfish decreases continuously with frozen storage days, mainly due to the formation of a large number of ice crystals in the tissue cells, which will squeeze the tissue structure, aggravating the physical damage of the tissue structure, and then causing the loss of juice in the tissue cells during the thawing process, while the water content of crayfish meat with chitosan nano-composite water-retaining agent in ultrasound treatment group was significantly higher than that in control groups, indicating that it may be due to the proper power of ultrasound treatment increased the phosphate ion of the water retaining agent into the muscle, as a consequence the pH of the muscle increased, and the hydrophilicity of actin enhanced, which inhibited the water loss from the muscles of crayfish meat.

3.2. Ice crystal size in crayfish meat

In general, ice crystals in tissues easily form during frozen meat products, and larger ones can damage the structure of the tissues. Therefore, the quality of frozen meat products largely depends on the morphology of the ice crystals formed [24]. Samples after repeated freeze–thaw cycles show significant differences in the morphology of ice crystals (Fig. 2a), especially the diameter and area of the ice crystals (Fig. 2b and c). With AUS and CED treatment, the average size of ice crystals was around 90.26 μm² and 113.73 μm², and the average diameter of ice crystals was 5.83 μm and 8.14 μm, respectively. The red areas correspond to muscle fibers and the white regions corresponded to cavities left by the ice crystals [25]. As the number of freeze–thaw cycles increased, the ice crystals became progressively larger, indicating poor tissue recovery and damage to the microstructure. After five freeze–thaw cycles, the mean area and diameter of the ice crystals increased significantly (p < 0.05). In contrast, the AUS sample showed less damage to the muscle fiber structure and produced smaller ice crystals, suggesting that the cavitation bubbles generated by ultrasound accelerated heat transfer and promoted nucleation and that the force generated by cavitation bubble explosions fractured large ice crystals into smaller ones [26]. Similarly, Sun et al. [27] found that ultrasound-assisted immersion freezing of common carp (Cyprinus carpio) with appropriate ultrasonic power treatment resulted in smaller and more uniform ice crystals, which helped maintain the integrity of the muscle tissue, as evidenced by small pores with high density, and they thought this may be attributed to cavitation bubbles produced by ultrasound-induced nucleation, and to microstreaming that may have fragmented pre-existing ice crystals into smaller nuclei to promote secondary nucleation, resulting in more ice nuclei and a shorter freezing time.

3.3. Physical properties of MPs

Solubility is an important indicator for protein denaturation or aggregation induced by freeze–thaw cycles [28]. Fig. 3a shows the effect of different treatments on the solubility of MPs in cooked crayfish after repeated freeze–thaw cycles. The solubility of MPs of CK was significantly higher than that of the other groups (p < 0.05). The solubility of CED samples reduced from 32.86 % to 18.29 % after 5 times of freeze–thaw cycles, suggesting that freeze–thaw cycles exacerbated the denaturation of the MPs. After repeated freezing and thawing, the WHC decreases, and the formation of larger ice crystals damaged the microstructure and exacerbated the extent of protein oxidation [24]. Thus, the structure of MPs was altered by repeated freeze–thaw cycles, which led to solubility change. Compared to other treatment groups, the solubility of MPs in AUS was the highest to be 30.26 % and that in CED was the lowest to be 18.29 %. As mentioned above, sonication can break large ice crystals formed during repeated freezing and thawing into small ice crystals, alleviating the degree of protein denaturation [29]. In addition, alginate oligosaccharide groups attached to the protein molecule can form many hydrogen bonds with water molecules and increase the charge attraction in protein systems and the electrical repulsion between protein molecules. A recent study indicated that a decrease in solubility might be attributed to the unfolding of the protein structure, exposure of hydrophobic residues, oxidation of reactive groups, and polymerization of proteins [30], suggesting the structural changes during freezing and thawing cycles (Fig. 3b and Fig. 4).

Fig. 3.

Changes in solubility (a), surface hydrophobicity (b), particle size (c) and zeta potential (d) of myofibrillar proteins on cooked crayfish subjected to different treatments. Different letters in the same column of the same indicator showed significant difference (p < 0.05).

Fig. 4.

Changes in Raman spectra (a), secondary structure content (b, 1: CK, 2, 3, 4: 0F-T; 5, 6, 7: 1F-T; 8, 9, 10: 2F-T; 11, 12, 13: 5F-T; each group was ordered as CED, AOS and AUS, respectively) and tertiary structure (c) of myofibrillar proteins on cooked crayfish subjected to different treatments.

Hydrophobic amino acid residues are usually found deep within the folded structure of the protein; therefore, surface hydrophobicity is commonly used as a criterion to assess conformational changes in proteins [31]. Fig. 3b shows the effect of different treatments on the surface hydrophobicity of MPs from cooked crayfish after repeated freeze–thaw cycles. After five freeze–thaw cycles, the surface hydrophobicity of CED, AOS, and AUS increased to 330.79 ± 5.21, 251.76 ± 12.19, and 175.19 ± 4.88, respectively, and it was found that the hydrophobicity of the AUS treatment group was significantly lower than that of the other treatment groups (p < 0.05). Liu et al. [32] found that ultrasonication may facilitate the tuna myofibrillar protein chains to expand on the surface, resulting in the exposure of hydrophobic amino acids and increasing the surface hydrophobicity. This differed from our results, mainly due to the high ultrasonic power used in their study (160 W for 6 min), which caused stronger cavitation. Moreover, the addition of alginate oligosaccharide may help to maintain the structure of MPs during ultrasonication, as indicated in Fig. 4.

Changes in the particle size of cooked crayfish MPs after different treatments are shown in Fig. 3c. The particle size increased significantly with the number of freeze–thaw cycles, indicating that repeated freeze–thaw cycles exacerbated protein denaturation and aggregation (p < 0.05), but the particle size in the AUS was significantly smaller than that of the CED group (p < 0.05). Similarly, Li et al. [22] showed that the formation of ice crystals disturbed the structure of proteins and exposed hydrophobic groups within pork MPs, thus leading to denaturation and aggregation of proteins, ultimately increasing the mean particle size. Wang et al. [33] investigated the combined multi-frequency ultrasound thawing of small yellowtail, where dual-frequency ultrasound pre-treatment resulted in smaller sizes of MPs particles.

The zeta potential of MP after different treatments are shown in Fig. 3d. As the number of freeze–thaw cycles increased, the absolute value of the zeta potential gradually decreased, indicating disruption of the structure of the protein molecules. It has been demonstrated that the higher the absolute value of the zeta potential, the greater the structural stability of MPs and the weaker the electrostatic repulsion between protein molecules, promoting aggregation and cross-linking between proteins and gradually increasing the average particle size of proteins [34]. In addition, the absolute values of zeta potential in the AUS treatment group were higher than those in the other treatment groups, indicating that ultrasonication could effectively delay the damage caused by repeated freeze–thaw cycles.

3.4. Spatial structure of crayfish protein

The secondary structure of cooked crayfish protein after different treatments determined via Raman spectra are shown in Fig. 4a. In general, the amide I band (1600–1700 cm⁻¹) is considered to be indicative of the secondary structure conformation of the protein, while the specific secondary structure (including α-helix, β-sheet, β-turn, and random coil) content was obtained by fitting and deconvolution [35]. As shown in Fig. 4b, there were significant differences in the secondary structures of the MPs obtained from CED, AOS and AUS treatment. The secondary structure of MPs in CK was mainly α-helix, which decreased after repeated freeze–thaw cycles, consistent with the results of Tan et al.[36]. High α-helix and β-sheets contents tend to tighten the protein structure, whereas a higher β-turn and random coil content represented a loose protein conformation [37]. After five freeze–thaw cycles, the α-helix and β-turn content of the AUS sample was 15.71 % and 8.35 % higher than that of CED, respectively, indicating that the secondary structure of MPs was more protected by AUS treatment. The α-helix content was negatively correlated with surface hydrophobicity (i.e., disruption of α-helices is accompanied by an increase in surface hydrophobicity), which is consistent with the results of surface hydrophobicity studies [33]. Qiu et al. [24] found that the α-helix content of frozen alfalfa muscle gradually decreased, while the proportion of β-sheets, β-turns, and random coils gradually increased as the frozen storage time increased owing to the gradual denaturation of MP, and repeated freeze–thaw cycles exacerbated protein oxidation, destabilizing the network of muscle fibers and the structure of MPs.

Tyrosine, tryptophan, and phenylalanine residues are very sensitive to changes in the surrounding microenvironment; therefore, the maximum fluorescence intensity (FI_max) reflects changes in the tertiary structure of the protein [38]. As shown in Fig. 4c, the fluorescence intensity of MPs gradually decreased with increasing freeze–thaw cycles. It has been shown that repeated freezing and thawing could induce conformational unfolding of MPs by disrupting intermolecular interactions, thereby exposing more hydrophobic groups, as evidenced by a decrease in fluorescence intensity [39]. The high fluorescence intensity of the AUS sample compared to that of the CED sample indicated that the structure of MPs was maintained to a greater extent. Wang et al. [40] used ultrasound (20 kHz, 500 W, intensity 30 %) to treat MPs and determined that it could improve the structural rigidity of myogenic fibronectin, preventing it from unfolding and altering its tertiary structure.

3.5. Gel electrophoresis

SDS-PAGE was used to analyze structural changes in MPs in crayfish meat, As shown in Fig. 5, three distinct bands corresponding to myosin heavy chain (MHC, 200 kDa), paramyosin (100 kDa), and actin (43 kDa) were obtained [35]. Compared with CK samples, the protein bands of the other samples changed significantly after repeated freeze–thaw cycles, with the protein bands becoming less pronounced and narrower, suggesting that repeated freeze-thawing exacerbated the degradation of MP. Li et al. [22] noted that, as the WHC of pork was significantly reduced during freezing and thawing, intracellular water spilled out, affecting the relative concentrations of solutes in the intracellular fluids and leading to protein denaturation. For each freeze-thawing cycle (most obvious in 5F-T), the bands in the AUS group were stronger than that in AOS, and the bands in AOS were stronger than those in CED, demonstrating that ultrasound-assisted penetration of alginate oligosaccharide into tissues could significantly minimize the changes in MPs. Wang et al. [33] found that the band density of MPs in small yellow croaker treated by dual-frequency ultrasonic thawing was closest to that of the fresh sample, indicating that there was less aggregation and degradation in the ultrasonicated samples.

Fig. 5.

SDS-PAGE patterns of myofibrillar proteins of cooked crayfish subjected to different treatments (1: Marker; 2: CK; 3, 4, 5: 0F-T; 6, 7, 8: 1F-T; 9, 10 11, 2F-T; 12, 13, 14: 5F-T. Each group was ordered as CED, AOS and AUS, respectively).

3.6. Microstructure of crayfish meat

The effect of ultrasound-assisted alginate oligosaccharide soaking on the microstructure of cooked crayfish after repeated freeze–thaw cycles was investigated using SEM, as shown in Fig. 6. Compared to the CK group, the structure of the muscle fibers changed significantly with increasing freeze–thaw cycles, indicating that the integrity of the muscle fibers was disrupted, the fibers became loose, and the gap between fibers gradually increased, which was an important factor indicating the significant decrease in the WHC of cooked crayfish after repeated freeze–thaw cycles [41]. However, ultrasound treatment allowed alginate oligosaccharide to enter the muscle fiber interstices and adsorb more water to the muscle surface, thereby increasing the WHC of cooked crayfish [42]. As can be seen from Fig. 2, the AUS sample exhibited less damage to the muscle fiber structure and produced smaller ice crystals during freeze–thaw cycles. Moreover, similar to our results, it has been reported that the cavitation effect can disrupt the structure of pork muscle fibers, thus reducing the resistance to external mass transfer and facilitating the diffusion of NaCl in pork muscle fibers [24].

Fig. 6.

Changes in microstructure of cooked crayfish subjected to different treatments (All the samples were enlarged with the magnification of 200×).

4. Conclusion

This study investigated the effect of ultrasound-assisted alginate oligosaccharide soaking (AUS) on the water retention of cooked crayfish during repeated freeze–thaw cycles. The utility of AUS treatment to improve water retention with 19.47 % higher and mobile water with 16.21 % higher than CED treatment after five freeze–thaw cycles. The average size of the ice crystals was around 90.26 μm² and average diameter of the ice crystals was 5.83 μm by AUS treatment, as well as the protein structure α-helix and β-turn content 15.71 % and 8.35 % higher than that of CED, demonstrating that AUS could reduce the mechanical damage of cooked crayfish caused by repeated freeze–thaw cycles, improve the quality of cooked crayfish, and contribute to the structural stability of MPs. Although ultrasound-assisted alginate oligosaccharide soaking harbors great potential for applications in improving the commercial value and shelf life of aquatic frozen products, the refrigeration (freezing rate and method) and sterilization (cold plasma or high pressure) are also important for frozen products and need to be considered in the further study. In this way, the freezing and preservation technology would be demanded to combine with ultrasound-assisted alginate oligosaccharide soaking treatment for better qualities of frozen products.

CRediT authorship contribution statement

Jiping Han: . Yingjie Sun: . Tao Zhang: Supervision. Cheng Wang: . Lingming Xiong: . Yanhong Ma: . Yongzhi Zhu: . Ruichang Gao: . Lin Wang: . Ning Jiang: Supervision.

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.

Appendix A. Supplementary data

The following are the Supplementary data to this article:

Data availability

Data will be made available on request.