Introduction

Aquaculture is now a critical food source and will likely be required to meet growing global food demands (Azra et al. 2021; Costello et al. 2020). With its high yields and low environmental footprints, bivalve aquaculture is considered one of the most environmentally sustainable ways to obtain animal protein (Bricker et al. 2018; Shumway et al. 2003; van der Schatte Olivier et al. 2020). Bivalve aquaculture, therefore, has contributed to a substantial amount of shellfish production in the past few decades and has become an essential seafood supply (Shumway et al. 2003). Among a variety of bivalve species, the endemic New Zealand green-lipped mussel, Perna canaliculus Gmelin 1791, has experienced increasing demand due to its high quality and nutritional value, now has an annual production worth NZ$394 million (Aquaculture New Zealand 2023).

Most of the green-lipped mussels produced are exported as processed frozen half-shells, making transporting this seafood product more logistically achievable than live mussels (Hickman 1991). As the frozen processes appear to diminish product freshness, and there is an increasing demand for higher quality mussels (Angane et al. 2020), live transport of mussel species has been investigated to improve its quality (Barrento and Powell 2016; Bernárdez and Pastoriza 2013; Tuckey et al. 2023; Zamora et al. 2019), like other seafood products such as crustaceans (Fotedar and Evans 2011). During live transportation, marine organisms, however, typically experience prolonged aerial exposure and/or closed seawater systems, where the environment is likely to deteriorate due to the accumulation of metabolic waste such as ammonia (Paterson et al. 2003). Furthermore, live transportation requires several mechanical processes, from harvesting to transport stages, which elevate mussel stress and mortality. Subsequently, this generates unnecessary waste due to mortality and product downgrading along the live seafood supply chain (Gould 1996; Stoner 2013; Woll and Bakke 2017). Exploration of new methods should, therefore, be prioritised to make the mussels more resilient to stress associated with transportation.

When energy supply cannot meet the increasing demand for combating stress, organisms may shut down non-essential and energy-consuming biological processes (e.g., protein synthesis), prioritising energy to deal with environmental stress that is likely to ensue (Hui et al. 2020; Sokolova et al. 2012). Such energy conservation strategies often result in a drop in overall metabolism due to reduced energy demand (Meinardus and Gäde 1981) reflected by depressed physiological processes, such as respiration and heart rate (HR) (Guppy and Withers 1999), which could also help to reduce generation and accumulation of metabolic waste in their surrounding environments. Depressed metabolism has been observed in various aquaculture species such as sea urchins, sea cucumbers, oysters, and cockles in response to changes in pH, oxygen level and food availability (Delorme and Sewell 2016; García-Esquivel et al. 2002; Huo et al. 2024) for suppressing metabolism of the green-lipped mussel, Perna canaliculus. The present study represents a key component of a wider programme of research aimed at enhancing live transportation outcomes.

Materials and methods

Mussel collection and maintenance

Adult mussels (shell length ~ 60 to 120 mm; n = 250) were collected from farms within Pelorus Sound (seawater temperature ~ 14 °C), South Island, New Zealand, in August 2022. The mussels were transported to the Cawthron Aquaculture Park (Nelson, New Zealand) and underwent biosecurity processing (as required by the New Zealand Government) to remove epibiota (mechanical shell cleaning followed by sequential 2-minute dips in freshwater to trigger complete valve closure, hypochlorite solution (200 ppm) and re-immersion in freshwater) within 24 h. Shell length (i.e., anterior–posterior distance) of all mussel individuals was obtained after experimentation using Vernier callipers (± 0.1 mm, SPI2000, Swiss). The mussels were then evenly distributed and maintained in four aquaria (150 L) kept at 15.6 ± 0.1 °C (i.e., ambient seawater temperature; mean ± SD) in flow-through filtered seawater (1 µm; salinity, 35) with algae supply (1:1 Tisochrysis lutea and Chaetoceros muelleri; 4.8 ± 0.7 µg Chl a L−1) for two weeks to recover from harvest and biosecurity handling processes. During this period, no mortality was observed, and the mussels started feeding immediately, develo** byssal attachments overnight.

Measurement of cardiac and ga** patterns of P. canaliculus

Mussels were exposed to different candidate suppressors—low temperature, low oxygen level and high concentration anaesthetic immersion (i.e., MgCl2·6H2O) known for reducing the metabolism of P. canaliculus (Cheng et al. 2024). For each suppressor, there were four experimental levels (one of them as control, see following sections and Table 1), and the mussels were exposed to each of them for 2 h. This time of exposure was selected based on previous research showing that this time was sufficient to allow any metabolic changes (Dunphy et al. 2015, 2018). During these exposures, mussels were selected for measurement of heartbeat and ga** (see the following section for the detail of measurements) at 30 minutes (T1) and 120 minutes (T2) after the exposure started.

Table 1 Summary of experimental conditions of (a) temperature, (b) oxygen levels and (c) increasing anaesthetic concentration treatments, where mussel heart rate and ga** behaviour were measured
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Temperature

Filtered seawater (1 µm; salinity, 35) was adjusted and maintained to different temperature regimes (i.e., 14 (control), 4, 6, and 8 °C; Table 1a) by chillers. Aerators and circulation pumps were set up to ensure the oxygen supply and homogeneity of temperature. The temperatures were recorded with Envloggers (27 mm diameter, ElectricBlue, Portugal). For each temperature, 16 mussels were evenly allocated into two 10-L tanks with temperature-controlled filtered seawater. Eight mussels were randomly sampled from the two tanks in each temperature treatment for each sampling time point to obtain their heartbeat and ga** traces simultaneously (i.e., ∑n = 4 temperatures × 2 time points × 8 mussels = 64 individuals (shell length, 81.5 ± 5.8 mm; mean ± SD)).

Heartbeat traces were measured using infrared pulse sensors (Vishay Semiconductors, CNY70; see Burnett et al. 2013) attached to the mussel shell near the dorsal posterior hinge. Ga** traces were recorded by attaching Hall sensors to the mussels. For each mussel, a magnet (~ 1 cm diameter) was attached to the posterior margin of one valve and a Hall sensor was attached to the opposing margin using a superglue. Both heartbeat and Hall sensors were connected to a PowerLab (16/35, ADInstruments, Australia) with 2 V and 10 V ranges, respectively, and a sampling rate of 1 ks−1. Heartbeat and ga** traces were visualised and recorded with LabChart™ 8 software (ADInstruments, Australia). The Hall sensor signal (voltage) changes proportionally to the magnetic field strength; the wider the gape, the smaller the signal due to the weaker magnetic field resulting from the increased distance between the magnet and the Hall sensor. After the sensors had been attached, the mussels were left undisturbed for 10 minutes prior to the measurement. Heartbeat and ga** traces were recorded for 20 minutes.

Heartbeat traces were processed with triangular smoothing with 301 samples to clean the noise off the signal. Heart rates (HR) were counted and calculated as beats per minute (bpm), excluding the unclassified heartbeat traces, to minimise the chance of misidentification of the heartbeat for heart rate calculation. The setting “standard integral” with timed reset every 60 s was applied to the heartbeat traces to calculate the area of heartbeat traces per minute. The pseudo-cardiac output (PCO; in an arbitrary volt-minute unit, au), which was calculated as the total area of the heartbeat traces for every minute, was obtained by extracting the maximum value of the integral per minute. Since the amplitude of the heartbeat traces changes with the displacement of the heart tissues of mussels during the cardiac cycle (Burnett et al. 2013), the total area of heartbeat traces per minute (i.e., PCO) can be a good proxy for the volume of fluid content passing through the atrial and ventricular chambers of a mussel as an indicator for cardiac activity level (Giomi and Pörtner 2013).

The raw ga** traces of the mussels were processed with a 1 Hz low pass filter in the Lab Chart 8 software. The processed ga** traces were down-sampled to 10 s−1 and subsequently exported for analysis. Such resolution has been shown to be sufficient to capture the fine valve movement of mussels (Robson et al. 2009). The signal of the mussels when they were completely sealed was marked and identified. Subsequently, 99th percentiles of such ga** traces were used to establish the fully closed value for the ga** data, which were transformed to relative ga** magnitude (RGM) for comparison of ga** patterns of mussel individuals under different experimental conditions using the equation below:

$$\frac{\text{Fully closed voltage}-\text{Current voltage}}{\text{Fully closed voltage}} \times 100\text{\%}=\text{RGM}$$

Dissolved oxygen level

Four treatments with different oxygen levels (i.e., 8 (control), 0.5, 1, and 3 mg O2 L−1; Table 1b) covering mild to severe hypoxic conditions, which were previously shown to depress mussel metabolism (Cheng et al. 2024; Diaz and Rosenberg 1995), were achieved by bubbling air or a mixture of air and nitrogen gas into filtered seawater (1 µm; salinity, 35); any temperature fluctuation (14.4 ± 0.2 °C) was monitored with a thermocouple (Lutron, TM947SD, Taiwan). During the experiment, the oxygen levels (i.e., dissolved oxygen, DO) were also monitored with an optical oxygen metre (Pyroscience, FirestingO2, Germany). For each oxygen level, 16 mussels were evenly allocated into two 10-L tanks with temperature-controlled filtered seawater. At each time point, eight mussels were used from each DO treatment (i.e., ∑n = 4 DO levels × 2 time points × 8 mussels = 64 individuals (shell length, 81.2 ± 6.6 mm)). Mussels were attached to the infrared pulse and Hall sensors for measuring HR and ga** patterns, respectively (see the “Temperature” section for the details of setup and data processing). Unfortunately, due to instrumental error, ga** traces of mussels, and subsequently relative ga** magnitude, at both time points of 1 and 3 mg O2 L−1, and the first time point of 8 mg O2 L−1 could not be obtained.

Anaesthetic

Aqueous solutions with different concentrations of the food-safe anaesthetic (EFSA-FAF et al. 2019), MgCl2·6H2O (i.e., 0 (control), 30, 40, and 50 g L−1), were achieved by the addition of MgCl2·6H2O (hereafter MgCl2) and a mixture of sodium chloride (NaCl), seawater, and freshwater to minimise the effects caused by variation in salinity and osmolality (Table 1c). Such concentrations have previously been shown to lower bivalve metabolism successfully (Cheng et al. 2024; Heasman et al. 1995; Suquet et al. 2009). Despite the fluctuation in osmolality of the solution, a previous study showed mussels were tolerant to short-term exposure (< 4 h) to change in osmolality (800–1000 mmol kg−1; McFarland et al. 2013). For each anaesthetic level, 16 mussels were evenly allocated into two 10-L tanks. At each time point, eight mussels from each anaesthetic level were attached to infrared pulse and Hall sensors for acquisition of heartbeat and ga** traces, respectively (i.e., ∑n = 4 anaesthetic concentrations × 2 time points × 8 mussels = 64 individuals (shell length, 81.6 ± 6.3 mm); see the “Temperature” section for the details of setup and data processing). All treatments were well-aerated throughout the experiment, and temperatures were monitored with a thermocouple and remained at 14.2 ± 0.4 °C.

Statistical analysis

The data for heart rate, pseudo-cardiac output and relative ga** magnitude obtained from mussels exposed to different conditions were analysed with two-way analysis of variance (ANOVA) using linear mixed-effects models from the R package “lmer” (Bates et al. 2015) to investigate the effects of different levels of selected inducers (i.e., temperature, oxygen or MgCl2; fixed factor) and exposure duration (i.e., two time points; fixed factor) and their interaction while taking into account the effects of inter-individual variance (i.e., individual; random factor). Post-hoc multiple comparisons were conducted using the R package “emmeans” (Lenth 2022) to detect the differences in heart rates, pseudo-cardiac output and relative ga** magnitude with Benjamini–Hochberg correction of p-values to reduce chances of committing Type I error (i.e., false positive).

The relationships between heart rate (HR), pseudo-cardiac output (PCO), and relative ga** magnitude (RGM) of mussels sampled at different conditions were analysed by non-parametric Spearman’s rank correlation test (i.e., the correlation coefficient ρ between − 1 and 1, indicating negative or positive correlations) since the HR, PCO and RGM data (i.e., calculated every minute for each individual) did not pass the normality test (Shapiro–Wilk test).

Results

Response to reduced temperature

Pseudo-cardiac output (PCO) and heart rates (HR) of the mussels were positively correlated (p < 0.001; Fig. 1a). Also, relative ga** magnitude (RGM) was positively associated with HR (p < 0.001; Fig. 1d) and PCO (p < 0.001; Fig. 1g). Temperature (p < 0.001) and exposure duration (i.e., time points; p < 0.01) had significant effects on HR of Perna canaliculus (Fig. 2a; Table 2a). HR dropped with temperature and increased with exposure duration. PCO, however, was not affected by temperature and exposure duration (p > 0.05; Fig. 2d; Table 2b). Temperature had a strong effect on mussels’ RGM (p < 0.01), which increased with temperature but remained stable between sampling time points (Fig. 2g; Table 2c).

Fig. 1
figure 1

Spearman’s rank correlations between pseudo-cardiac output and heart rate (a, b, and c), relative ga** magnitude and heart rate (d, e, and f), and relative ga** magnitude and pseudo-cardiac output (g, h, and i) of mussels from temperature (first column), low dissolved oxygen (second column), and MgCl2·6H2O (third column) treatments, respectively

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Fig. 2
figure 2

Boxplots for heart rates (a, b, and c), pseudo-cardiac output (d, e, and f) and relative ga** magnitude (g, h, and i) of mussel, Perna canaliculus, at different time points (i.e., T1, 30 minutes and T2, 120 minutes after the exposure), respectively, exposed various temperatures, dissolved oxygen levels, and MgCl2·6H2O concentrations. The solid line and blue square represent the median and the mean respectively (Con = control, i.e., for temperature (14 °C), low dissolved oxygen (8 mg O2 L−1), and MgCl2·6H2O (0 g L−1) treatments). Due to equipment failure, ga** data and subsequently relative ga** magnitude at both time points of 1 mg O2 L−1 and 3 mg O2 L−1, and the first sampling time point of 8 mg O2 L−1 could not be obtained

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Table 2 Results of two-way ANOVA to compare variation in (a) heart rates, (b) pseudo-cardiac output, and (c) relative ga** magnitude under different temperatures (Temp: control (i.e., Con = 14), 8, 6, and 4 °C) and time points (TP: T1 and T2)
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Response to reduced dissolved oxygen levels

There was a positive correlation between mussels’ PCO and HR (p < 0.001; Fig. 1b). RGM, in contrast, was not correlated to HR (p > 0.05; Fig. 1e) but positively correlated to PCO (p < 0.001; Fig. 1h). Mussels’ HR were interactively (p < 0.01) affected by dissolved oxygen level and exposure duration (i.e., time points; Fig. 2b; Table 3a). HR of the mussels at 1 mg O2 L−1, sampled at T1, had lower HR than those sampled at T2. Dissolved oxygen level and exposure duration did not affect mussels’ PCO (p > 0.05; Fig. 2e; Table 3b). There is also an increase in variation of HR and PCO with time for each condition, particularly, at 1 and 0.5 mg O2 L−1. RGM of mussels was impacted (p < 0.01) by dissolved oxygen levels where the mussels exposed to 0.5 mg O2 L−1 had higher RGM than those to 8 mg O2 L−1 (Fig. 2h; Table 3c).

Table 3 Two-way ANOVA to compare variation in (a) heart rates, (b) pseudo-cardiac output, and (c) relative ga** magnitude under different dissolved oxygen levels (DO: control (i.e., Con = 8), 3, 1 and 0.5 mg O2 L−1) and time points (TP: T1 and T2)
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Response to magnesium chloride

The PCO (p < 0.001; Fig. 1c) and RGM (p < 0.001; Fig. 1f) of mussels were positively correlated to HR, and RGM of the mussels was also positively correlated to PCO (p < 0.001; Fig. 1i). Mussel HR were affected by the interaction (p < 0.05) of MgCl2 concentration and exposure duration (i.e., time points; Fig. 2c; Table 4a). In the presence of MgCl2, mussel HR decreased with exposure duration. Mussels’ PCO was only affected by MgCl2 concentration (p < 0.01; Fig. 2f; Table 4b), remaining consistent between time points. Pairwise comparisons showed that the PCO of mussels without MgCl2 was higher than that of mussels exposed to MgCl2. Mussel RGM was only affected by MgCl2 concentration (p < 0.05; Fig. 2i; Table 4c) rather than the exposure time. The subsequent pairwise comparison, however, did not detect any differences in RGM among MgCl2 concentrations.

Table 4 Two-way ANOVA to compare variation in (a) heart rates, (b) pseudo-cardiac output, and (c) relative ga** magnitude under different MgCl2·6H2O concentrations (Conc: control (i.e., Con = 0), 30, 40 and 50 g L−1) and time points (TP: T1 and T2)
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Changes in cardiac activities and ga** patterns across conditions

Changes in heart rate, pseudo-cardiac output and relative ga** magnitude varied across conditions relative to “control” mussels (Table 5). HR of mussels at low temperatures could be depressed by 50–100%, whereas low dissolved oxygen (DO) levels only decreased mussels’ HR by 1.5–51%, while HR of mussels exposed to MgCl2 decreased by 36–97%. Mussels’ PCO in low temperatures decreased by 64–100%. There were, however, 7–350% increases in PCO of mussels in low DO levels. In the presence of MgCl2, mussels’ PCO decreased by 44–76%. Mussels in low temperatures had the lowest RGM (depressed by 27–88%), while mussels in low DO levels and the presence of MgCl2 had increased RGM by 115–191% and 11–152%, respectively.

Table 5 Changes (i.e., %) of heart rate (HR), pseudo-cardiac output (PCO), and relative ga** magnitude (RGM) of mussels, Perna canaliculus, measured at different time points (i.e., T1 and T2) exposed to various temperature, dissolved oxygen (DO), and anaesthetic (MgCl2·6H2O) levels relative to the ambient conditions (i.e., control)
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Discussion

Relationship between HR, PCO, and RGM

The present study utilised cardiac and behavioural biometrics to elucidate the efficacy of potential pre-transport treatments. Overall, a significant, positive relationship was found between the biometrics of relative ga** magnitude (RGM), heart rate (HR) and pseudo-cardiac output (PCO) of the green-lipped mussel, Perna canaliculus. Valve openness could regulate physiological processes through impacting oxygen supply and, in turn, the metabolic activities of bivalves (Davenport and Fletcher 1978; Trueman et al. 1973). For example, the mussel, Mytilus edulis, which was recovered in oxygenated seawater following an oxygen-deprivation environment, showed immediate valve ga** to increase oxygen supply, accompanied by increased HR (Coleman and Trueman 1971). In the present study, individuals of P. canaliculus with a more prominent gape could, therefore, indicate higher oxygen demand. Mussels with higher HR generally also had higher PCO, suggesting more haemolymph passing through the heart within a specific period. Such results, however, are not universal among ectothermic invertebrates (de Wachter and McMahon 1996; de Wachter and Wilkens 1996). For example, the increased HR of the green crab, Carcinus maenas, from 60 to 90 bpm did not correspond to an increase in the amount of haemolymph passing through the heart of the crab (Giomi and Pörtner 2013). Although PCO appears to be a helpful proxy for inferring cardiac activity, it only illustrates the relative rather than absolute changes in the amount of haemolymph passing through the heart of a mussel. There is a paucity of information on the simultaneous measurement of cardiac activities and valve movement on Perna canaliculus. The physiological and behavioural relationships present in this study, therefore, may serve as a pioneer dataset of P. canaliculus in different environments. The variability of cardio-behavioural responses may also help differentiate the mussel individuals with higher resilience to prolonged environmental stress, which could be used for selective breeding (Powell et al. 2017).

Mussel response to low temperature

The body temperature of ectothermic organisms, such as mussels, is dependent on the environmental temperature, which governs most temperature-dependent biochemical reactions (Arrhenius 1889) and, subsequently, metabolism and performance within thermal tolerance windows (e.g., Angilletta et al. 2002; Huey and Kingsolver 1989; Huey and Stevenson 1979; Wieser 1973). Decreased body temperature due to drop** environmental temperature could decrease enzyme activities and thus suppress metabolism. This can be reflected in depressed rates of physiological functions such as respiration and heart rate (see the review of Schulte 2015). Although HR and PCO were generally correlated in the present study, the significant drop in HR did not mirror the drop in PCO with decreased temperature. This was likely due to high inter-individual variability and irregular heart contraction (e.g., bradycardia) across temperatures, resulting in higher within-group (i.e., within each temperature) than between-group (i.e., between temperatures) variations. The reduced rates of physiological processes, such as heart rate, also imply decreased oxygen demand, which could be accompanied by reduced valve ga** (Tang and Riisgård 2016). As such, temperature could be a factor modulating the ga** patterns of bivalves (Comeau et al. 2012; Hernandis et al. 2018), which likely accounted for the general decrease in RGM of P. canaliculus with declining temperature. These responses, however, were likely dependent on the exposure duration, as mussels may subsequently enter a cold coma stage with their valves opened after prolonged exposure to cold temperatures, which has been shown in mussels Mytilus spp. (Jansen et al. 2007).

Mussel response to low oxygen

Perna canaliculus used in the present and preceding studies demonstrated high tolerance to an oxygen-deficient environment (Cheng et al. 2024), which has also been observed in congenerics Perna viridis and Perna perna which can survive prolonged exposure (four days to 4 weeks in seawater) of severe hypoxic and anoxic conditions (Hicks and McMahon 2005; Wang et al. 2011), which could be linked to their diverse energetic strategies during anaerobiosis (Gäde 1983a). When oxygen availability is limited, mussels could depress their aerobic metabolism to minimise energy demand, which can be observed in decreased physiological rates, such as respiration and heart rates, as shown in mussels Mytilus edulis and Perna perna (Hicks and McMahon 2002; Shick et al. 1986). Similarly, in the present study, P. canaliculus also showed depressed averaged HR after 30 minutes when held in severe hypoxia (i.e., 1 mg O2 L−1). Valve closure in response to acute hypoxia has been observed in other bivalves such as the oyster, Crassostrea virginica (Porter and Breitburg 2016). After an additional hour of exposure, mussels had increased gape, which may indicate an attempt at enhancing oxygen acquisition capacity due to increased irrigation of the mantle cavity (Jørgensen et al. 1988; Jørgensen 1990), resulting in increased metabolic rate, again illustrating the positive correlation between ga** magnitude and heart rate. There was a general decrease in HR and an increase in PCO at decreasing oxygen levels, which contradicted the positive correlation between HR and PCO. This could be driven by increased variation of HR and PCO of the mussels with time, particularly at severe hypoxia (i.e., 1 and 0.5 mg O2 L−1). Similar patterns were recorded in the scallop, Argopecten irradians, which had increased variance in HR at 1 mg O2 L−1 (Gurr et al. 2018). Due to this increased uncertainty associated with inter-individual response, hypoxia may not be an appropriate method to lower the metabolism of P. canaliculus.

Mussel response to MgCl2

When MgCl2 is applied, Mg2+ blocks Ca2+ channels and prevents Ca2+ from entering the cell. This suppresses neurotransmitter release (e.g., glutamate, acetylcholine; Namba et al. 1995; Sugi 1971; Szent-Györgyi 1975), thus resulting in failure of the posterior adductor muscle to contract results in valve opening. Such a situation explains the increased RGM of the mussels exposed to MgCl2, although such behavioural response was variable across MgCl2 concentrations. The relaxing effects of MgCl2 also limited the filtration activity and oxygen acquisition of the mussel, limiting the energy-generating processes of P. canaliculus (Azizan et al. 2021). As such, mussels may minimise energy demands by suppressing physiological processes, which could be indicated by decreased HR and corresponding PCO. The effect of MgCl2 on mussels was more prominent with increased exposure duration, and 2 h was sufficient to reduce mussels’ metabolism as evidenced by lower HR at T2 (~ 1 bpm) than at T1 (~ 5 bpm;), as longer exposure time presumably allowed more thorough diffusion of Mg2+ into mussel tissues. Exposing mussels to 40 g L−1 MgCl2 was the most effective way to reduce their metabolism, as individuals exposed to such conditions had almost 100% reduction in HR within 2 h. Conversely, a higher concentration of 50 g L−1 MgCl2 effectively relaxed oysters, Crassostrea gigas and Saccostrea glomerata, within 2–6 h (Butt et al. 2008; Suquet et al. 2009), suggesting taxa-specific MgCl2 dosage and exposure duration.

Scope of conditions to reduce the metabolism of mussels for live transport

Reducing stress through the suppression of metabolism has the potential to improve live transport of seafood to market (Lorenzo et al. 2020; Pozhoth and Jeffs 2022; Robertson et al. 2018). Low-temperature conditions have been commonly explored for both crustaceans and shelled molluscs live transport (e.g., Ben-Asher et al. 2020; Barrento and Powell 2016; Larssen et al. 2021; Tuckey et al. 2023; Zamora et al. 2019). Using the biomarkers of HR and PCO, we found that 4 °C was the most efficient temperature to depress the metabolism of P. canaliculus, as indicated by the apparent cessation of HR and PCO. Alternatively, pharmaceutical studies on live seafood transport mainly focus on crustaceans, such as crabs, lobsters and shrimps, which were demonstrated to successfully reduce mortality (5–10%) during live transport (e.g., Ben-Asher et al. 2020; Pozhoth and Jeffs 2022; Zhang and Li 2024). Such applications are, however, limited in molluscs, particularly, in mussels. In our study, the presence of 40 g L−1 MgCl2 reduced metabolism (97%) within 2 h and showed promise as a metabolic suppressant. Compared to other anaesthetics such as eugenol, benzocaine and MgSO4, MgCl2 solution is comparatively straightforward to prepare and is a more convenient and economical food-safe metabolic suppressant, and is, therefore, more commonly applied on aquaculture molluscs for various handling processes (Acosta-Salmón et al. 2005; Aquilina and Roberts 2000; Heasman et al. 1995; Suquet et al. 2009). Also, bivalves exposed to MgCl2 are less prone to spawn (Heasman et al. 1995), which is extremely important for the mussel market as the gonad fullness affects mussels’ quality and price. In contrast, mussels treated with oxygen-deficient conditions could not effectively reduce the metabolism as they did not show a strong indication of depressed metabolism (~ 50%) with high inter-individual variations. Such anoxic/hypoxic conditions could also bring more negative impacts to the mussels due to prevalent anaerobic metabolism (Gäde 1983b), which may affect their meat quality (Wells and Baldwin 1995).

Conclusions

The present study sought to identify the appropriate conditions to reduce the metabolism of the green-lipped mussel, Perna canaliculus, so it becomes inactive and insentive to the external environmental disturbance for live transport. Hypothermic conditions and immersion in food-safe MgCl2 solution both successfully reduced the metabolism of the mussel Perna canaliculus. Future work on the further comparison of both conditions on mussel metabolism, metabolite accumulation and their effects on mussel taste and texture will, therefore, be required to optimise the pre-treatment of green-lipped mussels for live transport. Further effort is needed to identify effective live transport approaches by simulating long-distance live transport processes in the air using the developed pre-treatment followed by recommendations to the stakeholders (e.g., the mussel suppliers and logistic providers) on applying such techniques in the live mussel supply chain.