Phytoplankton uptake and growth rate in the Japanese egg cockle Fulvia mutica
To clarify the relationship between the quantity of food ingested by and the growth rate of the Japanese egg cockle Fulvia mutica (Reeve), we conducted a laboratory breeding experiment for 2 weeks and estimated the chlorophyll a (chl-a) concentrations in water and the increments in shell length and soft-body weight of this species under five chl-a concentrations. Moreover, we compared the relationship between cockle growth (changes in soft-body weight and shell length) and their feeding environment observed in the laboratory experiment with the results of a field investigation conducted at two sites in the Sea of Japan, Kumihama Bay (35°38′5″ N, 134°54′00″ E) and Kunda Bay (35°33′30″ N, 135°15′4″ E). The changes in soft-body weight were similar in both laboratory and field investigations, but those in shell length were not. We, therefore, considered shell length changes as unsuitable for evaluating the relationship between growth and feeding in F. mutica. Based on the changes in soft-body weight, it was possible to classify the feeding environment of this species into the following three types: (1) < 1.52 μg chl-a L−1, negative feeding environment for cockle growth; (2) 1.52–5.71 μg chl-a L−1, neutral feeding environment for cockle growth; (3) > 5.71 μg chl-a L−1, positive feeding environment for cockle growth (growth increased with increasing chl-a concentration up to about 11 μg chl-a L−1). These results indicate that maintaining chl-a concentration in the breeding water within 5.71–11 μg chl-a L−1 is desirable for rearing Japanese egg cockle.
KeywordsFood demand Bivalve Mollusk Feeding environment Ingestion rate Clearance rate
The Japanese egg cockle, Fulvia mutica (Reeve), is a bivalve that inhabits the inner bays along the coast of Japan, excluding those in Hokkaido. This species is also widely distributed across the Korean Peninsula and the coast of China, and imported into Japan (Fujiwara 2009) where it is popular as shellfish sushi. Generally, marketed F. mutica are 3–4 g (stripped shellfish) and not treated as a high-class food material. In contrast, the F. mutica caught in Kyoto, Japan, are ten times larger (stripped shellfish weight 30–40 g) and their large, thick bodies are appreciated for their good taste. Therefore, F. mutica caught in Kyoto are often traded at high prices and have become special products (Fujiwara 2009).
Reports from several regions indicate large annual fluctuations in both catch and stock size of F. mutica (Tian and Shimizu 1998; Fujiwara 2009; Yang et al. 2011). Moreover, in Japan, the resources used by this species have collapsed in many areas (Tian and Shimizu 1997). In response, the Kyoto Prefectural Agriculture, Forestry, and Fisheries Technology Center started working on the development of seed production technology in 1976, aiming to ensure a stable supply of this species (Iwao et al. 1993). In addition, the full-scale F. mutica farming started by fishermen in 2000 has steadily increased its production since then (Tanimoto et al. 2015). However, in recent years, mass mortality of this organism occurred frequently during the culturing period, resulting in production declines. This mortality is likely related to recent climate changes which have resulted in higher water temperatures in summer, low salinity seawater due to heavy rain, generation of dysoxic water masses, and shortages of food. Currently, we are attempting to use aquaculture techniques to avoid mass mortality resulting from climate change. Specifically, a method for adjusting F. mutica to an appropriate water depth, without negatively influencing their survival and growth, has been developed and relies on monitoring the vertical profiles of several environmental factors in the aquaculture area. These factors include water temperature, salinity, dissolved oxygen concentration, and chlorophyll a (chl-a) concentration, which are used as indicators of the amount of phytoplankton available as food for the bivalves. Hourly measurements of these parameters are automatically performed by a lift-functioning conductivity, temperature, and depth (CTD) sensor installed in the aquaculture raft. To effectively operate such an aquaculture system, it is necessary to clarify the relationship between each environmental factor and the growth and/or survival of cockles. To date, the effects of water temperature, salinity, and dissolved oxygen concentration on the survival of other cockle species have been examined in detail in laboratory experiments (Nogami et al. 1981; Marsden and Bressington 2009; Tanimoto et al. 2015; Taylor et al. 2017; Peteiro et al. 2018). However, there is little information on the relationship between the feeding environment and the growth or survival of F. mutica and, as far as we know, no detailed studies examining these relationships have been conducted on this species in laboratory conditions.
The present study aimed to elucidate the relationship between the growth of F. mutica and its feeding environment (chl-a was used as an indicator of food availability). To achieve this aim, we examined the growth rate of this species in a 2-week laboratory rearing experiment under five different feeding conditions, i.e., different chl-a concentrations. Moreover, to ascertain to what extent the results of laboratory experiments represent those in the natural environment, we conducted field observations and compared them with the results of our laboratory experiments.
Materials and methods
The juvenile F. mutica used for the experiment were artificially produced in the Kyoto Prefectural Agriculture, Forestry, and Fisheries Technology Center and then farmed in Kunda Bay, Kyoto, Japan (35°33′30″ N, 135°15′4″ E) for about 2 months. Before the experiment, the individuals were kept in aerated, algae-free seawater that flowed through a breeding tank, for a period of 2 days. Shell length was 42.3 ± 2.4 mm (mean ± SD, n = 40) at the start of the experiment. All procedures performed in this study were in strict accordance with the “Guidelines for Animal Experimentation of Seikai National Fisheries Research Institute (SNFRI), Japan Fisheries Research and Education Agency, Japan” and “Guidelines for the Care and Use of Live Fish at SNFRI”.
Cultivation of the phytoplankton used for the feeding experiment in the laboratory
Components of the seawater medium used for Chaetoceros sp. culture
Sterilized filtered seawater
Vitamin mixture solutionb
bVitamin mixture solution
Feeding procedure in the laboratory experiment
Estimating phytoplankton ingestion quantity in the laboratory experiment
Estimating growth rates in the laboratory experiment
In the present study, growth rates were evaluated based on changes in the shell length and dry soft-body weight of juvenile cockles. The shell length of cockles in each breeding bottle was measured at the start and end of the laboratory experiment. Based on these measurements, the average daily extension rate (mm d−1) was calculated for each individual. To determine dry soft-body weight, the juvenile cockles were immediately dissected after the shell length measurement at the end of the breeding experiment, and their body tissues were dried to constant weight for 24–48 h at 60 °C. The soft-body weight of 15 cockles of the same lot as those used in the breeding experiment (n = 25) was also measured in the same manner to represent the initial dry weight (DW). Based on these measurements, the average daily weight increase rate (mg DW d−1) was calculated for each individual.
Estimating the relationship between the chl-a concentration in ambient water and the quantity of phytoplankton ingested by cockles
Field investigation site and estimation of F. mutica growth rate in situ
Field measurement of chl-a concentration
During the field study, we measured the chl-a concentration around the containers in which cockles were cultivated. At Kumihama Bay, vertical chl-a concentration profiles were automatically determined at every hour using the CTD data logger (DS5X; Hydrolab, Loveland, Colorado, USA). The daily chl-a concentration was taken as the average value of every 24 h. At Kunda Bay, the chl-a concentration at 6 m depth was measured every week using a portable CTD (MS5; Hydrolab).
Estimating the quantity of ingested phytoplankton in the field investigation
The quantity of phytoplankton ingested by cockles in the field study was estimated as follows. At each water depth at which the containers were suspended, the average chl-a concentration during the investigation period was calculated from the daily fluctuations (as described in the previous section), and these average values were regarded as the chl-a concentrations of the ambient water to which the cockles were exposed. The quantities of phytoplankton ingested by the field cockles were estimated based on the results of the laboratory experiment obtained for the clearance rate and on the average chl-a concentration values measured in the ambient water at the field sites.
Least squares linear regressions of cockle growth rate (soft-body weight and shell length) against daily quantity of ingested phytoplankton were performed. An F-distribution was used to calculate the significance of the slope and the 95% prediction intervals for the slope and intercept.
Laboratory feeding experiment
Summary of chlorophyll a (chl-a) concentrations in the breeding water used for each cockle and estimated quantity of ingested phytoplankton
Average chl-a concentrations in the breeding water of storage tanka (μg chl-a L−1)
Chl-a concentrations in breeding bottleb (μg chl-a L−1)
Total ingested quantity during the 2-week experiment (μg chl-a individual−1)
Daily ingested quantity (μg chl-a d−1 individual−1)
Relationship between the quantity of ingested phytoplankton and F. mutica growth
Based on these equations, the minimum amount of ingested phytoplankton required for cockle growth was 93.53 μg chl-a d−1 individual−1 (estimated chl-a concentration of ambient water: 1.52 μg chl-a L−1) for soft-body weight increase and 30.32 μg chl-a d−1 individual−1 (0.49 μg chl-a L−1) for shell length growth.
Chlorophyll a (chl-a) concentrations and cockle growth obtained in the field investigation in 2015
Average chl-a concentration (μg chl-a L−1)
Average dry soft-body weight (g)
Average shell length (mm)
Aug 7–Sep 29 (52 days)
7.83 ± 6.33 (n = 50)
0.15 ± 0.04 (n = 10)
1.70 ± 0.37 (n = 10)
26.8 ± 2.4 (n = 10)
54.0 ± 4.2 (n = 10)
Aug 7–Sep 29 (52 days)
5.23 ± 3.70 (n = 50)
0.18 ± 0.04 (n = 10)
1.12 ± 0.37 (n = 14)
29.5 ± 1.8 (n = 10)
50.7 ± 5.1 (n = 14)
Aug 27–Sep 29 (33 days)
4.24 ± 1.98 (n = 33)
0.98 ± 0.15 (n = 10)
1.67 ± 0.23 (n = 19)
49.7 ± 2.5 (n = 10)
59.6 ± 3.3 (n = 19)
Aug 27–Sep 29 (33 days)
4.83 ± 2.22 (n = 33)
1.43 ± 0.22 (n = 10)
1.49 ± 0.20 (n = 10)
50.5 ± 2.5 (n = 10)
56.6 ± 2.2 (n = 10)
Aug 27–Sep 29 (33 days)
5.39 ± 2.66 (n = 33)
1.11 ± 0.19 (n = 10)
1.68 ± 0.36 (n = 9)
46.5 ± 2.4 (n = 10)
58.0 ± 4.5 (n = 9)
Aug 20–Sep 15 (26 days)
1.47 ± 0.44 (n = 5)
0.67 ± 0.11 (n = 10)
0.66 ± 0.15 (n = 15)
43.5 ± 2.2 (n = 10)
44.1 ± 3.9 (n = 15)
Ambient water characteristics measured in the field investigation in 2015
Dissolved oxygen (mg L−1)
Aug 7–Sep 29 (52 days)
25.7 ± 2.4 (n = 50)
22.5 ± 7.4 (n = 50)
7.5 ± 1.2 (n = 50)
Aug 7–Sep 29 (52 days)
26.4 ± 1.8 (n = 50)
29.5 ± 2.4 (n = 50)
7.0 ± 1.0 (n = 50)
Aug 27–Sep 29 (33 days)
24.8 ± 1.0 (n = 33)
32.0 ± 0.4 (n = 33)
5.8 ± 0.5 (n = 33)
Aug 27–Sep 29 (33 days)
24.6 ± 0.8 (n = 33)
32.8 ± 0.2 (n = 33)
5.1 ± 0.8 (n = 33)
Aug 27–Sep 29 (33 days)
24.4 ± 0.6 (n = 33)
33.1 ± 0.1 (n = 33)
4.5 ± 1.0 (n = 33)
Aug 20–Sep 15 (26 days)
25.6 ± 1.6 (n = 5)
33.7 ± 0.6 (n = 5)
6.5 ± 0.2 (n = 5)
In the aquaculture industry, it is important for fishery products to grow fast and systematically. In other words, it is necessary to understand the relationship between food consumption and growth. This is especially true for bivalves, as this basic information is necessary when considering sea area selection or production density. However, when examining the growth of bivalves, two indicators are generally used: body weight increment and shell size growth (shell length, shell height, and shell width) (e.g., Franz 1993; Alunno-Bruscia et al. 2001; Larson et al. 2014). In the present study, we examined the growth of F. mutica based on soft-body weight and shell length increments, and compared the results of the laboratory experiment with that of the field investigation. Although for the relationship between the soft-body weight increment and feed consumption, the measurements from the field investigation fell almost entirely within the range of the 95% prediction interval of the regression analysis based on the laboratory experiment (Fig. 9), for the increment of shell length, the measurements from the field investigation were remarkably different from that of the laboratory experiment (Fig. 10). Many studies have reported that the increase in bivalves’ soft-body weight and shell length are affected by various factors in their habitat (Malone and Dodd 1967; Seed 1976; Bayne and Worrall 1980; Kautsky 1982; Franz 1993; Alunno-Bruscia et al. 2001; Wong and Levinton 2004; Berge et al. 2006; Hiebenthal et al. 2012). In particular, it has been reported that food quantity affects the growth of bivalves differently when considering soft-body weight increase or shell-size increase (Franz 1993; Alunno-Bruscia et al. 2001). Franz (1993) reported that the mussel Geukensia demissa conserves shell growth at the expense of body weight increase in the case of food depletion. Using Mytilus edulis, Alunno-Bruscia et al. (2001) demonstrated that the effects of reducing or ceasing soft-body weight increase due to food depletion were stronger than the effects of reducing shell growth. However, the authors pointed out that the materials used for the production of bivalve shells and soft tissues partly originated from different sources. It is known that the shell is formed largely through the deposition of ions, mostly calcium from seawater (Wilbur and Saleuddin 1983), and that the organic matter content of the shell is below 5% (Jørgensen 1976; Price et al. 1976). In addition, many studies have reported that shell growth still occurs in starved or undernourished mollusks (e.g. Orton 1925; Palmer 1981; Lewis and Cerrato 1997). Thus, bivalves may not necessarily require nutritional supplementation by feeding to form shells, while nutrient supply via feeding seems indispensable for soft-body growth as it contains many organic components. Still, habitat conditions (e.g. crowding due to population density, properties of sediments, etc.) directly and physically affect bivalve shell formation (Seed 1968; Brown et al. 1976; Newell and Hidu 1982). Although we cannot ascertain the cause, in the present research, the differences in shell length increment between the laboratory experiment and the field investigation might have been influenced by factors other than the food quantity ingested. In summary, it is considered that the best indicator to directly evaluate the influence of the quantity of ingested food on bivalve growth would be to estimate the increment in soft-body weight. Furthermore, because aquaculture is a protein-providing industry, information on efficiently increasing bivalve soft-body weight is important. Thus, the results of the present study are discussed below focusing on the increment of soft-body weight.
In the present study, the relationship between ingested phytoplankton quantity and soft-body weight increment was represented by a linear regression (Fig. 7). According to this equation, the amount of ingested food required for F. mutica soft-body weight increase was at least 93.53 μg chl-a d−1 individual−1 (estimated chl-a concentration in ambient water: 1.52 μg chl-a L−1). When a similar ingested food quantity (estimated chl-a concentration in ambient water) was calculated from the lower limit of the 95% prediction interval, the amount of ingested food required for F. mutica soft-body weight increase was 351.44 μg chl-a d−1 individual−1 (5.71 μg chl-a L−1) (Fig. 7). Up to about 700 μg chl-a d−1 individual−1 (11.37 μg chl-a L−1), which was the highest quantity of food ingested under the experimental conditions, the soft-body weight of F. mutica increased as the amount of food increased. Based on these results, it is possible to divide the feeding environment for breeding F. mutica into three categories according to ambient chl-a concentration: (1) < 1.52 μg chl-a L−1, negative feeding environment for cockle growth; (2) 1.52–5.71 μg chl-a L−1, neutral feeding environment for cockle growth (individuals either grow or not); (3) > 5.71 μg chl-a L−1, positive feeding environment for cockle growth (soft-body weight increases with increasing chl-a concentrations up to about 11 μg chl-a L−1). Saito et al. (2007) reported the relationship between the resource characteristics of Ruditapes philippinarum and their habitat environment, based on field surveys. The authors divided the habitat chl-a concentrations into four stages [(1): bad—(4): good] depending on the clams’ resource situation: (1) < 1 μg chl-a L−1, (2) 2–4 μg chl-a L−1, (3) 4–6 μg chl-a L−1, (4) ≥ 6 μg chl-a L−1. Comparing the results obtained here with that of Saito et al. (2007), and although the studied species and research methods were different, a similar tendency can be recognized regarding the classification of chl-a concentrations in the ambient water for bivalve growth. In the future, it will be necessary to clarify the relationship between bivalves and their feeding environment by examining and comparing results for other bivalve species. Notably, however, our laboratory experiments were carried out at 25 °C, in filtered seawater with salinity 33 psu, and normoxic conditions. It is known that the filtration rate of bivalves is influenced by factors such as temperature, salinity, and dissolved oxygen concentration of the ambient water (e.g. Sará et al. 2008; Enríquez-Ocaña et al. 2012; Nieves-Soto et al. 2013; Riisgård et al. 2013; Kang et al. 2016; Tang and Riisgård 2018). Because bivalves are filter feeders, changes in filtration rates are directly related to changes in the quantities of ingested food. In addition, temperature and salinity also affect the metabolism of bivalves, and therefore change in these factors result in change in the growth rate of bivalves (e.g. Liu et al. 2018; Zhang et al. 2018; Sanders et al. 2018; Haider et al. 2019); for example, low water temperatures might reduce energy requirements, and as a result, less food would be required for growth; under high water temperature, the opposite phenomenon might be observed. So far, the biological effects of several environmental factors, such as ambient water temperature, salinity, and dissolved oxygen concentration, on F. mutica have been investigated in the laboratory (Nogami et al. 1981; Tanimoto et al. 2015). Regarding the influence of water temperature, the oxygen consumption of cockles increased with increasing water temperature in the range of 15–24 °C, and reached a maximum value of around 25 °C, before decreasing gradually from 26 to 28 °C (Nogami et al. 1981). In addition, it was reported that survival rate on the 5th day of breeding would be about 60% in normal seawater (salinity 34 psu) at 28 °C (Tanimoto et al. 2015). With respect to salinity, it was reported that salinity levels which would not affect the survival of cockles were 22 psu or more, in water temperatures of 20–26 °C (Tanimoto et al., 2015). As for the influence of dissolved oxygen concentration, under conditions of less than 2 mL L−1, decrease in oxygen consumption, the appearance of abnormal open-shell or mortality individuals were observed (Nogami et al. 1981). Based on the present results, the laboratory experimental conditions used here are desirable conditions for the high physiological activity of F. mutica. Thus, depending on the conditions of the ambient water (including temperature, salinity, and dissolved oxygen), the minimum food consumption required for cockle growth (and the minimum chl-a concentration required in the ambient water to ensure minimal food intake) could be overestimated using the regression line obtained in the present study.
It is known that bivalves change their filtration rates according to food-particle concentrations (Winter 1978; Navarro and Winter 1982), and highly concentrated feeding environments, in particular, inhibit bivalve filtration activities (Toba and Miyama 1993; Pérez-Camacho et al. 1994; Nagasoe et al. 2011). In addition, under high food-particle concentrations, bivalves filter a quantity of food particles exceeding the amount that could be ingested per unit time, and excess particles must be excreted as pseudofeces (Widdows et al. 1979). Because, as mentioned above, a highly concentrated food-particle environment might cause bivalve feeding inhibition, it has been emphasized that maintaining food-particle concentration within an appropriate range is necessary for efficient bivalve rearing (Thompson and Bayne 1974; Toba and Miyama 1993). Thus, determining food concentration thresholds that inhibit bivalve feeding activity and growth is as important as grasping the minimum amount of food required for growth. Unfortunately, the phytoplankton concentrations within the cockle-breeding bottles used in the present study were all equally low, despite the chl-a concentration gradient generated in the water of storage tanks. Consequently, we could not clarify the food concentration threshold that would inhibit cockle growth. Widdows et al. (1979) mentioned that the amount of material filtered per hour would be equivalent to the maximum ingestion rate at the threshold concentration of pseudofeces production (before any rejection occurs). This could be interpreted to imply that feeding inhibition occurs above the threshold concentration at which bivalves excrete pseudofeces. The results of the laboratory experiment performed here revealed an average chl-a concentration of about 11 μg chl-a L−1 as the upper limit of food concentration (Figs. 6, 7) above which no excretion of pseudofeces was observed. Thus, food concentrations above 11 μg chl-a L−1 inhibit the growth of F. mutica, but these should be clarified by detailed experimentation. To examine the growth of F. mutica under a highly concentrated feeding environment using the experimental system of the present study (Fig. 1), it will be necessary to adjust chl-a concentrations in the water of stock tanks to considerably higher levels than used here; in other words, it will be necessary to keep a certain amount of food in the ambient water inside the breeding bottles even after bivalve filtration.
The comparison of the results of the laboratory experiment and field investigation implies that soft-body weight increment is a better evaluation index than shell length increment to illustrate the effects of bivalve feeding on growth. In addition, the feeding environment (chl-a concentration in ambient water) could be classified into three stages based on the soft-body weight increment of cockles: (1) < 1.52 μg chl-a L−1, negative feeding environment for cockle growth; (2) 1.52–5.71 μg chl-a L−1, neutral feeding environment for cockle growth (individuals either grow or not); (3) > 5.71 μg chl-a L−1, positive feeding environment for cockle growth (growth increases with increasing chl-a concentration up to about 11 μg chl-a L−1). For the reasons stated above, it would be desirable to cultivate F. mutica at chl-a concentrations between 5.71 and 11 μg chl-a L−1. The information obtained in the present study can contribute to the efficient breeding and farming of F. mutica and in seedling production; for example, in the selection of sea areas and depths with suitable food environments, in the estimation of breeding cockle density to maintain a suitable food environment, and in the consideration of feed volumes for indoor breeding. However, we could not observe growth inhibition of F. mutica due to high food concentration, which must be clarified by further detailed experiments in the near future.
This study was part of the results obtained with the support of the NARO Bio-oriented Technology Research Advancement Institution [Grant number 14526938]. We kindly thank K. Tarutani, Y. Matsuyama, T. Kurihara and the anonymous referees for their constructive comments, which have improved our manuscript.
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