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Maturation process of broodstock of the pen shell Atrina pectinata (Linnaeus, 1767) in suspension culture.

ABSTRACT The processes of gametogenesis and maturation in broodstock of the scaly-type pen shell Atrina pectinata in suspension culture were studied throughout the year. Gametogenesis in both sexes was initiated in February, when the water temperature was lowest. Gonad development and subsequent maturation occurred from March to September as the water temperature and food density increased. Spent gonads were found in October. Analysis of oocyte diameter frequencies in females just after spawning suggested that pen shell females spawn repeatedly during the breeding period. A number (1.6%) of hermaphroditic animals were also identified. Glycogen contents gradually decreased during the vitellogenic and mature stages. These results suggested that gonad development of pen shells in suspension culture was characterized by annual synchronicity in males and females, a spawning period from May to September resulting from monocyclic gametogenesis throughout the year, and an inverse relationship between gonad development and glycogen level. This study provides basic information applicable to mariculture of pen shells for commercial production.

KEY WORDS: Atrina pectina, pen shell, broodstock, gametogenesis, reproductive cycle, suspension culture


Pen shell, Atrina pectinata (Linnaeus, 1767) belonging to the family Pinnidae is a large (shell length up to 30 cm), suspension-feeding bivalve common along the coasts of Japan, Korea, and China (Okutani 1997). It is an infaunal bivalve found in muddy to sandy sediment habitats, from tidal flats to shallow subtidal environs to 20 m depth (Yurimoto et al. 2003). The two types of pen shell--scaly type and smooth type--are defined by the presence or absence of squamation of the shell surface (Torigoe 1985). Morphological comparison and isozymic analyses have revealed that these two types should be taxonomically distinguished (Yokogawa 1996). In southwestern Japan, both types are the most commercially important taxa. The fishery operates mainly in Ariake Bay on Kyushu Island, and a total wet weight per year of more than 30,000 t of pen shell was harvested in the 1960s. However, since 1992 production in this area has decreased rapidly, totaling as little as 37 t in 1999 (Kawahara & Ito 2003, Yurimoto et al. 2003). In addition, mass mortalities occurred in most major fishing grounds in 2003 and 2004 (Maeno et al. 2006, Yurimoto et al. 2008b), with a devastating impact on the pen shell fishery. The cause of this sharp recent reduction in the standing stock of the pen shell is unknown but might have been caused by an increase in sediment load (Ito 2006), the predation by the bullnose ray (Kawahara et al. 2004), or viral infection (Maeno et al. 2006).

For the recovery of stock abundance, one effective short-term measure is to establish technical procedures that can be used in aquaculture. In aquaculture of bivalves, one of the most critical elements is a regular supply of seed. This is necessary for successful artificial spat collection from broodstock to acquire knowledge of population characteristics with regard to gametogenesis, the reproductive cycle and spawning patterns (Gosling 2003). In an attempt to restore the stock abundance of the pen shells, efforts are currently under way to establish new techniques for seed production and thus prevent the reduction of natural stocks. In 2006, although the large-scale production of scaly-type pen shell larvae and juveniles was successfully developed in indoor tanks for the first time (Ohashi et al. 2008), the rates of fertilization and hatching, and the survival rate after settlement, were not stable. One of the influences on these phenomena could be gamete quality, which reflects the status of broodstock before spawning (i.e., gametogenesis and final maturation). However, the reproductive cycle of the pen shell has been examined only in populations in natural beds (Watanabe 1938, Yurimoto et al. 2005, Chung et al. 2006), and no information is available on the reproductive characteristics of the pen shell under artificial rearing conditions. Therefore, an understanding of the gametogenesis and maturation processes would provide us with essential information for artificial seed production and subsequently intermediate culture of the animal. Our objective was to examine gonad development, especially in terms of the gametogenesis and maturation processes of the pen shell (scaly-type), through histological observations of broodstock in suspension culture.


Environmental Parameters

Water temperature, salinity, and cell density of diatoms were recorded in situ monthly at a depth of 2 m at the suspension site.


More than 200 pen shell broodstock (shell length 141-221 mm) were collected by hookah diving at about 10-m depth in January 2007 from natural stock in the Seto Inland Sea off Kagawa Prefecture. They were immediately transported to the Seikai National Fisheries Research Institute at Nagasaki Prefecture and were reared in suspension at Mie Port in Nagasaki Prefecture from January 2007 to February 2008. Ten pen shells were sampled monthly and their shell length and shell height measured. The gonads were fixed in 10% seawater formalin for histology. Additionally, two hermaphroditic animals were found in March and June 2007, and the gonads of these were fixed for histological observation. The adductors muscle were frozen, and then stored at -40[degrees]C until used for glycogen content analysis.


Gonads of pen shells were examined histologically to determine the profiles of gametogenesis and the reproductive cycle. The mid part of the gonads was fixed in 10% seawater formalin for 24 h and then dehydrated in an alcohol series and embedded in paraffin wax. The paraffin blocks were sectioned to 4-[micro]m thickness and routinely stained with Mayer hematoxylin and eosin (H & E).

Distribution of Oocyte Diameters Just after Spawning

In the course of the investigation, spawning of two female pen shells was found in May and June 2007. The ovaries of these pen shells were excised just after spawning, and then fixed in 10% seawater formalin and processed as earlier mentioned for histological examination. The histological slides were observed under a microscope at x40 magnification. To ensure that each section passed through the center of the gamete, measurements were made only in oocytes that displayed a well-defined germinal vesicle. This operation was performed on 200 randomly chosen oocytes per individual.

Glycogen Contents

Glycogen content of soft tissues was determined from April 2007 to January 2008 by the anthrone method (Kamada & Hamada 1985). A piece of the adductor muscle (0.5-1.0 g) was removed and suspended in 1.5 mL of 30% KOH, then saponified by boiling to 100[degrees]C for 20 ruin. After the boiling, 2 mL of 90% ethanol and 0.25 mL of saturated sodium sulfate were added to the sample solution to extract glycogen. Glycogen in the sample was centrifuged (x3,000 g for 5 min.), and diluted with deionized water. Each subsample was added to a one-fifth volume of anthrone-sulfuric acid solution and boiled to 100[degrees]C for 15 min. After cooling, absorbance of the resulting colored complex was measured at a wavelength of 620 nm.




Environmental Parameters

We analyzed the monthly water temperature and salinity at the time of sample collection (Fig. 1). A seasonal change in water temperature was observed, peaking from August to September 2007 (28.3[degrees]C to 30.5[degrees]C) and decreasing gradually until its lowest point, winter (14.8[degrees]C in February 2007 and 15.7[degrees]C in January 2008). Salinity remained stable throughout the year (32.6-34.6 PSU), with the exception of a decrease in July 2007 (30.1 PSU) caused by heavy rainfall. Diatom cell densities showed a clear seasonal pattern characterized by three unequally-sized peaks (Fig. 2). Peaks were seen in April (5.7 x [l0.sup.5] cells/L), September (5.5 x [10.sup.5] cells/L), and October (4.1 x [l0.sup.s] cells/L) 2007. Cell densities remained low level from December to February (7.5 x [10.sup.4] cells/L to 0.9 x [10.sup.4] cells/L).


The gonad of the pen shell is not a discrete organ but is attached to the digestive gland (Fig. 3). Macroscopic observation between April and August revealed that the male gonads were whitish and the female gonads orange to red. Because of the limited nature of this observation period, gonad color was not a reliable indicator of gonad development or sex.




The testis and ovary were composed of several follicles surrounded by connective tissue. In males, examination of nuclear characteristics and cell size revealed that the germ cells could be classified into four stages: spermatogonium, spermatocyte, spermatid, and spermatozoa. The spermatogonia were attached to the follicular wall. The cells were 6-8 [micro]m in diameter and spherical or oval, with a nuclear diameter of about 5 [micro]m (Fig. 4A). The cytoplasm was stained lightly with hematoxylin, and the nucleolus was distinguishable within the nucleus. The spermatocytes were almost round and 3-4 [micro]m in diameter (Fig. 4A). The nucleus was stained with hematoxylin, and was 2 [micro]m in diameter. The spermatids were smaller than the spermatocytes and were located close to the lumen (Fig. 4B); their nuclei were spherical and about 1 1.5 [micro]m in diameter. The spermatozoa were spherical and l [micro]m in diameter. The heads of the spermatozoa were separated from the trabeculae and aligned in rows (Fig. 4B). The chromatin was small and completely condensed.


In females, early-stage oocytes were attached to the follicular wall and mature oocytes were located in the lumen. Analysis on the basis of cell size and morphology revealed five stages of female germ cell: the oogonium and four stages of oocyte. The oogonium was round and 10 12 [micro]m in diameter and the cells were attached to the inner side of the follicular wall (Fig. 5A). The previtellogenic oocyte was 10-20 [micro]m in diameter, and the nucleus was round and 10 [micro]m in diameter (Fig. 5A). The nuclei contained chromatin, which was dispersed throughout the nucleus, and the cells were attached to the follicular wall. The early vitellogenic oocyte was 25-30 [micro]m in diameter, with a nuclear diameter of 12 15 [micro]m (Fig. 5B). The cells were still attached to the follicular wall. The late vitellogenic oocyte was larger (30-65 [micro]m in diameter), with a nuclear diameter of 35-40 [micro]m (Fig. 5C); these cells were round or polygonal. Numerous yolk granules were dispersed throughout the cytoplasm and the nucleolus was clearly visible. The late vitellogenic oocytes were completely detached from the follicular wall. Matured oocytes before spawning had numerous yolk granules and no apparent nucleus: that is, germinal vesicle breakdown was observed (Fig. 5D). The diameter of mature oocytes ranged from 65-80 [micro]m.

Maturation Process

The process of maturation of the broodstocks of the pen shells was examined by histological observation. It was graded into 5 stages according to a scale of maturity (modified from Yurimoto et al. 2005): (1) resting, (2) developing, (3) mature, (4) partial spawning, and (5) spent.

Resting Stage

This stage was characterized by folded follicular walls, no traces of gametes, and the presence of connective tissue on the formed follicular walls (Fig. 6); therefore, the sex of the pen shell was unidentifiable. Individuals in the resting stage were found from December to February (Fig. 7).


Developing Stage

The gonad was characterized by cxpansion of the follicular walls and the appearance of spermatogonia along the walls (Fig. 8A). Primary spermatocytes proliferated rapidly and moved to the center of the lumen. In the late period of the developing stage, the size of the lumen increased and spermatids were sometimes present in the lumen, but not abundant. The developing stage of the pen shell occurred from February to May (Fig. 7).

Mature Stage

The spermatids and spermatozoa occupied most of the area of the lumen, and arranged themselves in radial rows toward the center of the lumen (Fig. 8B). The mature stage was observed from April to August (Fig. 7).

Partial Spawning Stage

Spermatozoa were released into the water, and the gonad still contained spermatids and spermatozoa in the lumen. The follicular wall was wrinkled and partly collapsed (Fig. 8C). The partial spawning stage was observed from May to September (Fig. 7).



Spent Stage

Mature spermatozoa were completely discharged and the follicular wall was almost empty (Fig. 8D). A few residual spermatozoa were seen around the wall. The wall had become wrinkled or degenerated and the gonad was obviously smaller. The spent stage was first observed in September; all shellfish examined had spent gonads in October, and the gonads continued to be spent until December (Fig. 7).


Developing Stage

The ovary consisted of a capsule-like structure, which contained oogonia and oocytes attached to the trabeculae (Fig. 9A). The developing stage was observed from February to June (Fig. 7).

Mature Stage

In the mature ovary, there were abundant late oocytes and matured oocytes. Most oocytes were polygonal (Fig. 9B). The nucleus had sometimes disappeared owing to germinal vesicle breakdown. The mature stage in the female occurred from May to August (Fig. 7).


Partial Spawning Stage

The follicles of the ovary appeared partly empty, indicating that mature oocytes had been released (Fig. 9C). The follicles contained mostly late vitellogenic oocytes and matured oocytes. Partial spawning occurred from August to September (Fig. 7).

Spent Stage

Mature oocytes were completely discharged and the follicles were almost empty (Fig. 9D). The follicular wall had become folded or degenerated and the gonads had decreased in size. A few residual oocytes were seen in the wall. The spent stage of the ovary occurred from August to December (Fig. 7).


Distribution of Oocyte Diameters Just after Spawning

Very few mature oocytes (diameter 65-80 [micro]m) were retained, and the most representative of oocyte diameters fell in the range 30-65 [micro]m (late vitellogenic oocyte) in both of the individuals examined (Fig. 10 & Fig. 11).



Two hermaphroditic animals found in March and June 2007 were passing through the same developmental stages as non-hermaphroditic individuals (Fig. 12A, B). The gonad stage of the sex components in the two individuals was synchronized (i.e., the one examined in March was in the developing stage and the one examined in June was in the mature stage).

Glycogen Contents

We examined the monthly change in glycogen content of adductor muscle (Fig. 13). The mean glycogen content peaked in April (37 mg/g), and then decreased gradually from April to July 2007. It then stayed low (between 8 and 22 mg/g) from July to December 2007. It increased again to 23 mg/g in January 2008.



Maturation Process

Histological observations of the gonads of pen shell broodstock in suspension culture revealed that gonad development of males and females was annually synchronous. Gametogenesis of both sexes commenced in February, and gonad development and maturation occurred from March to September. Completely spent gonads were found in October. These results suggested that the process of maturation of scaly-type pen shell broodstock was characterized by a unimodal cycle, and that the breeding period of this animal in suspension culture was from May to September. As a population, pen shells in suspension culture underwent an annual reproductive cycle that involved successive periods of gonadal differentiation, development, maturation, and spawning. The process of maturation of pen shell broodstock in suspension culture was similar to that of pen shells in natural beds (Yurimoto et al. 2005, Chung et al. 2006).

Gametogenesis in Relation to External Factors

The reproductive cycle of a species is a genetically controlled response to the environment, and the pattern of the reproductive cycle in a species is apparently determined through the coordination of successive reproductive events with changes in external factors (Sastry 1979). The successive events in the reproductive cycle may be affected differently by various kinds of factors, and a detailed analysis requires a separate evaluation of each phase. In pen shell broodstock in suspension culture, gamete differentiation commenced when the water temperature was lowest (14.8[degrees]C), and gametogenesis and maturation then coincided with increasing temperature from March to September. In pen shells in natural beds, although the initiation of gonad development occurred in February (Yurimoto et al. 2005), the water temperature was much lower (< 10[degrees]C) (Fuchigami et al. 2001). The difference in the water temperature initiating gamete differentiation at the two sites suggested that the triggering stimulus for initiation of gametogenesis was not water temperature but other external factors such as day length or food availability. No relationship was seen between salinity and gonad condition.


Spawning Characteristics

Quantitative analysis of oocyte diameter frequency seems to be a good quantitative descriptor of the maturational status in other bivalves (Laruelle et al. 1994, Lango-Reynoso et al. 2000, Gribben et al. 2004). We found two or three peaks in the distribution of oocyte diameter just after spawning in females, which retained a large number of vitellogenic oocytes. These facts showed that there was successive maturation of oocytes and asynchronous restoration of the gonad after spawning in the pen shell. These observations suggested that pen shell females spawned more than once during the breeding period.

Glycogen and Food Availability in Gametogenesis

Food availability to bivalve populations has been directly measured in terms of chlorophyll-a (Broom & Mason 1978, MacDonald & Thompson 1985), suspended particle matter (Yurimoto et al. 2008a), and energy content of seston (MacDonald & Thompson 1985). Gametogenesis in marine bivalves occurs at the expense of recently ingested food and/ or energy stored in various tissues. In bivalves, gametogenesis is an energy-demanding process, because the mobilization of nutrients to the gonad is essential for gamete development (Barber & Blake 1990), for which the main energy reserve is glycogen (Gosling 2003). The main storage tissue differs with the species: for example, it is the mantle in the mussel Mytilus edulis (Linnaeus, 1758) (Bayne et al. 1982), the midgut gland in the oyster Crassostrea gigas (Thunberg, 1793) (Yamamura & Watanabe 1964, Mori et al. 1965) and the adductor muscle in the scallop Patinopecten yessoensis (Jay, 1857) (Miyazono & Nakano 2000) and pen shell (Yurimoto et al. 2005).


In pen shells on natural beds, glycogen content is negatively correlated with the gonad development and is a useful parameter for characterizing physiological condition (Yurimoto et al. 2005). Similarly, here we found an inverse relationship between gonad development and glycogen content, suggesting that carbohydrates played an important role as energy sources for gametogenesis in pen shells in suspension culture and on natural beds. Gonad differentiation of the pen shell in suspension culture occurred when food was not abundant; however, the period of gonad development coincided with abundance in food. These observations suggest that adequate food supply is not imperative for gonad differentiation, whereas gonad development and maturation are closely coupled to food availability.

Our observations of the reproductive characteristics of pen shell broodstock should provide a valuable insight into the biology of this species and are crucial for broodstock management and seed production as well as for intermediate culture. The pen shell was recently recognized as a prime candidate for aquaculture in Ariake Bay, Japan, because it meets many of the biological and economic criteria necessary for candidate species. However, further applied studies of reproduction are required if we are to develop an economically viable protocol.


The authors thank Dr M. Minagawa of the Fisheries Research Agency, Seikai National Research Institute, Japan for his critical comments on the manuscript. Major funding for this study was provided by the Ministry of Agriculture, Forestry and Fisheries of Japan as part of a research project on the utilization of advanced technologies in agriculture, forestry and fisheries.


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(1) Seikai National Fisheries Research Institute, Fisheries Research Agency, 1551-8 Taira, Nagasaki, Nagasaki 851-2213, Japan; (2) Nagasaki Prefectural lnstitute of Fisheries, 1551-4 Taira, Nagasaki, Nagasaki 851-2213, Japan; (3) Institute for Environment and Biology, World Data Bank Co. Ltd., 1-6 Tonomui, Minami, Tokushima 779-2307, Japan

* Corresponding author. E-mail:

([dagger]) Present address: Japan International Research Center for Agricultural Sciences, 1-1 Ohwashi, Tsukuba, Ibaraki 305-8686, Japan
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Author:Maeno, Yukio; Suzuki, Kengo; Yurimoto, Tatsuya; Fuseya, Reiko; Kiyomoto, Setsuo; Ohashi, Satoshi; On
Publication:Journal of Shellfish Research
Article Type:Report
Geographic Code:9JAPA
Date:Aug 1, 2009
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