OOCYTE ATRESIA CHARACTERISTICS AND EFFECT ON REPRODUCTIVE EFFORT OF MANILA CLAM TAPES PHILIPPINARUM (ADAMS AND REEVE, 1850).
Understanding the dynamics of animal populations requires a firm knowledge of biological processes, particularly reproduction (Caddy 1989, Knights 2012, Costa et al. 2013, Delgado et al. 2013, Maunder & Deriso 2013). Considerable knowledge concerning the reproductive cycle of exploited marine bivalves has been accumulated since the early studies of the 1930s, documenting gametogenesis and spawning in many species of economic interest (see Lucas 1965, Sastry 1979, Mackie 1984, Gosling 2015 for reviews and references). Chief among these species is the Manila clam Tapes philippinarum (family Veneridae), the top-ranking marine aquaculture species worldwide (over 4 million tons of aquaculture production in 2014), with a total value three times greater than that of the familiar Pacific oyster Crassotrea gigas (FAO 2016). Various aspects of the reproductive cycle of T. philippinarum have been studied (Adachi 1979, Mann 1979, Beninger & Lucas 1984, Rodriguez-Moscoso et al. 1992, Robert et al. 1993, Laruelle et al. 1994, Xie & Burnel 1994, Chung et al. 2001, Park & Choi 2004, Delgado & Camacho 2007, Dang et al. 2010, Uddin et al. 2010, Uddin et al. 2012, Baek et al. 2014, Milani et al. 2017); however, the phenomenon of oocyte atresia has not been investigated. Although this form of oocyte degeneration has been mentioned and/or described in mature and residual oocytes in Pectinidae (Tang 1941, Christiansen & Oliver 1971, Dorange & Le Pennec 1989, Motavkine & Varaksine 1989, Le Pennec et al. 1991, Vaschenko et al. 1997, Borzone et al. 2003, Cantillanez et al. 2005, Beninger & Le Pennec 2006), Ostreidae (Lango-Reynoso et al. 2000, Dutertre et al. 2009), Mytilidae (Pipe 1987, Motavkine & Varaksine 1989, Suarez et al. 2005, Alonso et al. 2007) and Pinnidae (De Gaulejac et al. 1995), it seems to be an under-reported phenomenon in bivalve reproductive cycles in general (Beninger 2017). Oocyte atresia has been identified within the Veneridae, although its histological characteristics have not been described (Morvan & Ansell 1988, Meneghetti et al. 2004, Drummond et al. 2006, Casas & Villalba 2012).
The present study documents the phenomenon of atresia in the reproductive cycle of Tapes philippinarum. Two data sets were used, a detailed 4- and 5-mo study during the gametogenic period and a longer time series (26 mo) with fewer individuals. In addition to the qualitative description of this process, we use quantitative histological techniques to estimate its impact on reproductive effort ([R.sub.e]).
MATERIALS AND METHODS
Species, Sites, and Sampling
For reasons unclear to most workers, Tapes philippinarum has a relatively long list of competing generic names, of which Ruditapes is the most frequent in recent years. The reasons for selecting Tapes are outlined in Beninger and Boldina (2014).
The detailed histological study was carried out at a Tapes philippinarum and Cerastoderma edule culture operation, situated in an extensive mudflat aquaculture region on the French Atlantic coast. Twenty-three adult clams were haphazardly sampled every 2 wk (May to August 2015 and April to September 2016; 2.6-5 cm along the anteroposterior axis).
For the longitudinal time series, one fished and one unfished site were chosen on the French Atlantic coast: a recreational mudflat fishing site for Tapes philippinarum in the Gois passage, and an isolated, unfished site accessible only by boat; all three sampling sites were within 50 km of each other (Fig. 1). The sediment characteristics, water temperature, salinity, turbidity, and tidal regimes of the two sites were very similar (Boldina & Beninger 2013, Beninger & Boldina 2014, Boldina et al. 2014). In the course of other, unrelated manipulations, nearly commercial-size clams ([greater than or equal to]3 cm) were haphazardly sampled monthly at low tide from the unfished and fished sites (July 2010 to September 2012). The sampling took place at the incoming tide, often limiting the number of individuals sampled ([less than or equal to]5); in addition, several technical problems reduced the number of sampling dates. The number of females investigated is specified in Figures 5-7.
Sampled clams were separated from their shells and fixed on-site in ice-cold aqueous Bouin's solution for at least 48 h. Three-to 4-mm-thick slices were removed along the dorsoventral axis of the foot, continuing up to the dorsal extremity of the visceral mass. Samples were then rinsed overnight under running tap water, dehydrated in an ascending ethanol-Roti-Histol series, and embedded in paraffin. Sections were cut at 7 [micro]m and stained with a modified Masson's trichrome protocol (Martoja & Martoja-Pierson 1967, Beninger et al. 2010) using trioxyhematein (3 min), acid fuschine (2 min), orange G-phosphomolybdic acid (3 min), and fast green (1 min). Observations and analyses of the photomicrographs were performed using an Olympus Provis light microscope, and LUCIA GF 4.80 image capture and processing software. A total of 12 micrographs were archived for each female at the unexploited and fished sites, and nine micrographs for the farmed site, for later examination and stereological counts.
Preliminary investigations showed that the Tapes philippinarum gonad satisfied the requirements for stereological analysis: constant gonad tissue anatomical localization, synchronous gametogenesis throughout the gonad, sufficiently homogeneous gonad tissue, and sufficiently large patches of gonad tissue were available to perform counts (Beninger & Boldina 2012a). For the purposes of this study, only the oocyte types were quantified using stereological counts: atresic oocytes (AO), immature healthy oocytes (IO), and mature healthy oocytes (MO) (Beninger 1987, Beninger et al. 2001, Valdizan 2011; Fig. 2A).
Generally, three counts were performed on each of three sections per individual, and the data were pooled to provide mean counts for each cell type for each individual, date, and site. Counts were performed on all females sampled over the 26-mo study period at the unexploited and fished sites (July 2010 to September 2012) and over the gametogenic period at the farmed site (May to August 2015 and April to September 2016). A 13 x 14 counting grid was used for the unexploited and fished site micrographs, whereas an 11 x 11 grid was used for the farmed site. Given the small number of females for some samples, the range was used as an indicator of dispersion about the mean (Beninger & Boldina 2012b).
Qualitative Characteristics of Atresia
Using the modified Masson's trichrome staining protocol, healthy oocytes were readily identified with a pink cytoplasm and a rounded, well-defined nucleus; depending on the plane of section, a distinct nucleolus was also visible (Fig. 2). Healthy oocytes were either mature (mature healthy oocytes--regular rounded shape, separated from the acinal wall, Fig. 2B) or immature (immature healthy oocytes--pear shape, attached to the acinal wall with a peduncle, Fig. 2C).
Oocyte atresia in clams was characterized by the appearance of several characteristics, concomitantly or not. These characteristics were observed in both mature and immature oocytes; some affected the nucleus, others the cytoplasm.
Characteristics affecting the cytoplasm were as follows:
(1) Cytoplasmic discoloration (Fig. 3A-C). In most cases, the atresic cytoplasm stained more intensely, becoming darker compared with that of healthy oocytes. More rarely, the cytoplasm turned from a uniform pink-red to green and purple.
(2) Cytoplasmic retraction and detachment from the cell membrane (Fig. 3D-F, I). This was visible on histological slides as a clear space between the membrane and the cytoplasm, either on a small portion of the cell or involving most of the cell (Fig. 3D-F, I).
(3) An irregular geometric shape, noticeably different from the spherical shape of healthy oocytes (Fig. 3G-I).
Characteristics affecting the nucleus were as follows:
(1) Disappearance of the nucleoli (Fig. 3A-F, H-L). This characteristic is not sufficient on its own, because the section plane can pass above or below a nucleolus. In many cases, however, nucleoli are often histologically visible in actively synthesizing cells; thus their absence is a clue to atresia, especially when observed in a large number of oocytes, where it can indicate widespread atresia.
(2) Homogeneous chromatin, seen as a very uniform nucleus color, with no chromatin clumping (Figs. 3J-L and 4).
(3) Disappearance of the nucleus in medially sectioned cells (Fig. 3C, I).
The aforementioned characteristics may be more or less pronounced based on the chronological development of atresia. It is important to note that oocytes presenting different characteristics can be physically close to one another and also close to apparently healthy oocytes in the same acinus (Fig. 3D, H, J, K). Some parts of the clam gonad may be almost exclusively occupied by AO, whereas the rest of the gonad may be much less affected. In the overwhelming majority of observations, areas most severely affected were located in the dorsalmost region of the visceral mass.
Seasonal Pattern of Atresia
Histological observations allowed the identification of three major phases of the Tapes philippinarum oogenic cycle (Table 1).
Characteristics of atresia were observed in early and late vitellogenic oocytes (atresic immature oocytes), and in mature oocytes (atresic mature oocytes), beginning in April and throughout gametogenesis and postspawning resorption (Fig. 4). In July (Fig. 4B), oocytes were predominantly late vitellogenic or mature, and characteristics of atresia were again noted in both stages. During the resorption phase (September to November), most oocytes showed at least one characteristic of atresia, notably the disappearance of nucleoli (Fig. 4C), indicating generalized atresia in the gonad.
Quantification of Atresia
To assess the importance of oocyte atresia throughout the reproductive cycle of Tapes philippinarum, the following oocyte volume fractions were calculated (Weibel et al. 1966):
(1) Atresic volume fraction (AVF): the number of grid points occupied by AO divided by the total number of grid points occupied by all oocyte types.
AVF = [AO/IO + AO + MO]* 100
(2) Mature volume fraction (MVF): the number of grid points occupied by mature oocytes divided by the total number of grid points occupied by all oocyte types.
MVF = [MO/IO + AO + MO]* 100
(3) Immature volume fraction (IVF): the number of grid points occupied by immature oocytes divided by the total number of grid points occupied by all oocyte types.
IVF = [IO/IO + AO + MO]* 100
(4) Minimum atresic impact (MAI): the minimum impact of atresia on the oocyte population, expressed as
MAI = [AVF/AVF + MVF]* 100
The MAI was based on three assumptions: (1) MVF (composed of healthy, mature oocytes) represented the oocyte volume fraction with a high probability of being spawned as healthy; (2) AVF represented the oocyte volume fraction with no probability of being spawned as healthy oocytes; the fates of MVF and AVF were therefore known; (3) The fate of IVF was unknown as it could either remain healthy or become atresic. This index, therefore, represents the minimum oocyte volume fraction known to be atresic, compared with the total oocyte volume fraction whose fate is known.
The evolution of MVF and IVF values over the sampling period at the farmed site reveals several gametogenic cycles and spawns (Fig. 5). The AVF was at least 15% before the first spawn, increasing to 25%-30% in subsequent spawns. At the end of the reproductive period (September 2016), all residual oocytes were obviously destined for atresia. These results at the farmed sampling site in 2015 and 2016 extend and confirm the longer term observations from the fished and unfished sites (2010 to 2012; Figs. 6 and 7).
To clearly summarize these data, only those dates corresponding to MVF [greater than or equal to] 20% were chosen to represent the period of active gametogenesis (Table 2). The excluded periods were thus as follows:
(1) The end of the gametogenic periods, when most oocytes were atresic and undergoing resorption.
(2) The resting phase, when there was no gametogenesis.
(3) The beginning of the gametogenic periods, when most oocytes were in early gamete stages and their fate is thus unknown.
Over the active gametogenic period at all sites, approximately 33% of gamete volume was occupied by AO and the MAI was 45% (Table 2). No marked differences in volume fractions were noted between sites.
Histological Features of Atresia
Histological observations of the present study establish the following indicators of atresia: absence of the nucleolus (an early indicator), as well as nuclear degradation (irregular nuclear envelope or chromatin degradation), cytoplasmic discoloration and retraction, and cellular distortion (puzzle shape). Although it is not possible to establish a firm chronological sequence at this point, certain atresic characteristics do present a temporal sequence, especially for the nucleus (Fig. 8).
The features described previously correspond to the categories of characteristics previously outlined in the Bivalvia (Beninger 2017). No discernable differences were noted between pre- and postspawning atresia, indicating a common process.
Generalization of atresia within acini has been reported for several bivalve species, being easily recognized by major distortions of oocyte shape (Dorange & Le Pennec 1989, Suarez et al. 2005, Beninger & Le Pennec 2006, Dutertre et al. 2009). Such generalization may also occur in Tapes philippinarum acini, but this appears to lack synchrony, and cellular distortions are much less severe. Instead, clarification of the nucleus is the revealing feature.
Temporal Dynamics of Oocyte Atresia
Based on all of the data from the different study sites and years, the temporal dynamics of oocyte atresia in Tapes philippinarum are summarized in Figure 9. Oocyte atresia steadily increased in the spring, before the first spawning. A precipitous decrease in all oocyte types characterized the first spawning; from this point onward, throughout the subsequent gametogenic activity, both pre- and postspawning AO were present in the gonad simultaneously.
Of the three functional types of atresia proposed by Motavkine and Varaksine (1989), it is clear that the resorption phase of the reproductive cycle corresponds to residual atresia. The atresia observed before the resorption phase may be either physiological (i.e., a regulatory mechanism) or ecological (i.e., a response to unfavorable environmental conditions). Much further research will be necessary to refine this analysis.
Impact of Atresia on [R.sub.e]
To the authors' knowledge, this is the first study to quantify oocyte atresia and its effect on reproductive effort [R.sub.e]. Previously, Morvan and Ansell (1988) calculated a percent of AO in Tapes rhomboides, using a complex estimation based on oocyte diameters and the Williams' equation (Williams 1981); however, these authors only appear to have included mature AO in their estimations. The losses of 11 % fecundity in spring and 3.3% in summer reported by these authors, therefore, represent considerable underestimations.
The stereological technique used in this study is based on the number of counting points occupied by particular cell types. This type of data does not allow precise oocyte numbers to be determined, but it can obviously be used as a proxy for such numbers, in addition to representing the amount of energy invested. If all oocytes are viable, the [R.sub.e] simply equals the total volume of oocytes in the gonad at time If some oocytes are not viable, the effective reproductive effort (E[R.sub.e]) is the total oocyte volume fraction minus the volume fraction of all AO:
E[R.sub.e] = [R.sub.e] - AO
The results of the present study show that the minimum level of atresia in the Tapes philippinarum reproductive cycle was 15%, at the beginning of gametogenesis; this volume fraction climbed to approximately 33% during active gametogenesis, when both pre- and postspawning atresia are present (Figs. 9 and 10). Furthermore, the MAI obtained during active gametogenesis, from the three sites, was approximately 45%. Thus, nearly half of the oocyte volume fraction produced during active gametogenesis would he lost to atresia, reducing [R.sub.e] by approximately 45%. It should be remembered that this is the MAI, so the figure may well surpass 50%. As there was no evidence of AO resorption during active gametogenesis (no empty cells or macrophage invasion), it is assumed that these cells are lost at spawning; atresia, therefore, seems to represent a net loss of energy for T. philippinarum females.
It is useful to place this result in the context of [R.sub.e] estimates, approximated by indices such as the condition index (Lucas & Beninger 1985). To illustrate this point, such a condition index was calculated at the farmed site for 18 Tapes philippinarum individuals concomitantly sampled biweekly over 20 mo of one study period (March 2015 to December 2016, Fig. 11). The high values of this index during active gametogenesis are obviously misleading and should therefore be interpreted with caution because almost half of the gamete volume produced was atresic. The data of the present study show that although tissue: shell weight condition indices can be used to quantify reproductive investment, they cannot be used to indicate reproductive outcome.
From the preceding, it is clear that atresia can be a major cause of oocyte mortality in Tapes philippinarum; in itself, this result assists in the understanding of the high-level mortalities typically found in the early life stages of this and many other bivalve species, as well as allowing more realistic estimations of [R.sub.e] and fecundity.
The authors thank M. Christophe Hery, president of the Professional Clam Fishers Association of Vendee, for assistance at the unfished site, as well as Pascal Chellet, vice-president of the Regional Professional Shellfish Farmer Commission, for his interest and support for this project. Student interns Marie Petitguyot, Baptiste Serandour, and Tanguy Moreau assisted with histological preparation. This work was financed by the EPAT program of the Conseil Regional des Pays de la Loire, contract 2015 02488.
Adachi, K. 1979. Seasonal changes of the protein level in the adductor muscle of the clam, Tapes philippinarum (Adams and Reeve) with reference to the reproductive seasons. Comp. Biochem. Physiol. 64A:85-89.
Alonso, P. S., C. A. Gonzalez, P. M. Garcia & F. S. J. Serrano. 2007. Atresia gonadal durante el ciclo gametogenico de Mytilus galloprovincialis Lamarck, 1819 cultivado en la ria de Vigo (noroeste de la peninsula Iberica). Bol. Inst. Esp. Oceanogr. 23:3-10.
Baek, M. J., Y. J. Lee, K. S. Choi, W. C. Lee, H. J. Park, J. H. Kwak & C. K. Kang. 2014. Physiological disturbance of the Manila clam, Ruditapes philippinarum, by altered environmental conditions in a tidal flat on the west coast of Korea. Mar. Pollut. Bull. 78:137-145.
Beninger, P. G. 1987. A qualitative and quantitative study of the reproductive cycle of the giant scallop, Placopecten magellanicus, in the Bay of Fundy (New Brunswick, Canada). Can. J. Zool. 65:495-498.
Beninger, P. G. 2017. Caveat observator: the many faces of prespawning atresia in marine bivalve reproductive cycles. Mar. Biol. 164:163.
Beninger, P. G. & I. Boldina. 2012a. Rapport du projet IMPAP. Etude sur l'impact de la peche a pied: une approche multidisciplinaire, vol. 2: 2011-12.
Beninger, P. G. & I. Boldina. 2012b. Strengthening statistical usage in marine ecology. J. Exp. Mar. Biol. Ecol. 426-427:97-108.
Beninger, P. G. & I. Boldina. 2014. Fine-scale spatial distribution of the temperate infaunal bivalve Tapes (=Ruditapes) philippinarum (Adams and Reeve) on fished and unfished intertidal mudflats. J. Exp. Mar. Biol. Ecol. 457:128-134.
Beninger, P. G., R. Cannuel, J. L. Blin, S. Pien & O. Richard. 2001. Reproductive characteristics of the archaeogastropod Megathura crenulata. J. Shellfish Res. 20:301-307.
Beninger, P. G. & M. Le Pennec. 2006. Structure and function in scallops. In: Shumway, S. E. & G. J. Parsons, editors. Scallops: biology, ecology and aquaculture, 2nd edition. Amsterdam, The Netherlands: Elsevier Science Publishers, pp. 123-227.
Beninger, P. G. & A. Lucas. 1984. Seasonal variations in condition, reproductive activity, and gross biochemical composition of two species of adult clam reared in a common habitat: Tapes decussates L. (Jeffreys) and Tapes philippinarum (Adams & Reeve). J. Exp. Mar. Biol. Ecol. 79:19-37.
Beninger, P. G., A. Valdizan, P. Decottignies & B. Cognie. 2010. Field reproductive dynamics of the invasive slipper limpet, Crepidula fornicata. J. Exp. Mar. Biol. Ecol. 390:179-187.
Boldina, I. & P. G. Beninger. 2013. Fine-scale spatial structure of the exploited infaunal bivalve Cerastoderma edule on the French Atlantic coast. J. Sea Res. 76:193-200.
Boldina, I., P. G. Beninger & M. Le Coz. 2014. Effect of long-term mechanical perturbation on intertidal soft-bottom meiofaunal community spatial structure. J. Sea Res. 85:85-91.
Borzone, C. A., P. R. Pezzuto & Y. A. G. Tavares. 2003. Caracteristicas histologicas del ciclo reproductivo de Euvola ziczac (Linnaeus) (Pectinidae Bivalvia del littoral sur-sudeste del Brasil). Rev. Bras. Zool. 20:763-772.
Caddy, J. F. 1989. Marine invertebrate fisheries: their assessment and management. New York, NY: John Wiley & Sons. 752 pp.
Cantillanez, M., M. Avendano, G. Thouzeau & M. Le Pennec. 2005. Reproductive cycle of Argopecten purpuratus (Bivalvia: Pectinidae) in La Rinconada marine reserve (Antofagasta, Chile): response to environmental effects of El Nino and La Nina. Aquaculture 246:181-195.
Casas, S. M. & A. Villalba. 2012. Study of perkinsosis in the grooved carpet shell clam Ruditapes decussata in Galicia (NW Spain). III. The effect of Perkinsus olseni infection on clam reproduction. Aquaculture 356-357:40-47.
Christiansen, H. E. & S. R. Oliver. 1971. Sobre el hermaphrodismo de [much less than] Chlamys teheulcha [much greater than] d'Orb. 1846 (Pelecypoda, Filibranchia, Pectinidae). [On the hermaphroditism of the sea scallop d'Orb. 1846 (Pelecypoda, Filibranchia, Pectinidae)]. An. Soc. Cient. Argent. 191:115-127.
Chung, E. Y., S. B. Hur, Y. B. Hur & J. S. Lee. 2001. Gonadal maturation and artificial spawning of the Manila clam Ruditapes philippinarum (Pelecypoda: Veneridae), in Komso Bay, Korea. J. Fish. Sci. Technol. 4:208-218.
Costa, P. M., S. Carreira, M. H. Costa & S. Caeiro. 2013. Development of histopathological indices in a commercial marine bivalve (Ruditapes decussatus) to determine environmental quality. Aquat. Toxicol. 126:442-454.
Dang, C., X. De Montaudouin, M. Gam, C. Paroissin, N. Bru & N. Caill-Milly. 2010. The Manila clam population in Arcachon Bay (SW France): can it be kept sustainable? J. Sea Res. 63:108-118.
De Gaulejac, B., M. Henry & N. Vicente. 1995. An ultrastructural study of gametogenesis of the marine bivalve Pinna nohilis (Linnaeus 1758) I. Oogenesis. J. Mollus. Stud. 61:375-392.
Delgado, M. & A. P. Camacho. 2007. Influence of temperature on gonadal development of Ruditapes philippinarum (Adams and Reeve, 1850) with special reference to ingested food and energy balance. Aquaculture 264:398-407.
Delgado, M., L. Silva & A. Juarez. 2013. Aspects of reproduction of striped venus Chamalea gallina in the Gulf of Cadiz (SW Spain): implications for fishery management. Fish. Res. 146:86-95.
Dorange, G. & M. Le Pennec. 1989. Ultrastructural study of oogenesis and oocytic degeneration in Pecten maximus from the Bay of St. Brieuc. Mar. Biol. 103:339-348.
Drummond, L., M. Mulcahy & S. Culloty. 2006. The reproductive biology of the Manila clam, Ruditapes philippinarum, from the north-west of Ireland. Aquaculture 254:326-340.
Dutertre, M., P. G. Beninger, L. Barille, M. Papin, P. Rosa, A.-L. Barille & J. Haure. 2009. Temperature and seston quantity and quality effect on field reproduction of farmed oysters, Crassostrea gigas, in Bourgneuf Bay, France. Aquat. Living Resour. 22:319-329.
FAO. 2016. FAO yearbook. Fishery and aquaculture statistics. 2014. Rome, Italy. 30 pp.
Gosling, E. 2015. Reproduction, settlement and recruitment. In: Marine bivalve molluscs. Oxford: Fishing News Books, pp. 157-202.
Knights, A. M. 2012. Spatial variation in body size and reproductive condition of subtidal mussels: considerations for sustainable management. Fish. Res. 113:45-54.
Lango-Reynoso, F., J. Chavez-Villalba, J.-C. Cochard & M. Le Pennec. 2000. Oocyte size, a means to evaluate the gametogenic development of the Pacific oyster, Crassostrea gigas (Thunberg). Aquaculture 190:183-199.
Laruelle, F. J., J. Guillou & Y. M. Paulet. 1994. Reproductive pattern of the clams, Ruditapes decussatus and Ruditpes philippinarum on intertidal flats in Brittany. J. Mar. Biol. Assoc. U.K. 74:351-366.
Le Pennec, M., P. G. Beninger, G. Dorange & Y.-M. Paulet. 1991. Trophic sources and pathaways to the developping gametes Pecten maximus (Bivalvia: Pectinidae). J. Mar. Biol. Assoc. U.K. 71:451-463.
Lucas, A. 1965. Recherches sur la sexualite des mollusques bivalves. These de Doctorat, Universite de Caen.
Lucas, A. & P. G. Beninger. 1985. The use of physiological condition indices in marine Bivalve aquaculture. Aquaculture 44:187-200.
Mackie, G. L. 1984. Bivalves. In: Tompa, A. S., N. H. Verdonk & J. A. M. Van den Biggelaar, editors. The Mollusca, vol. 7: reproduction. Orlando, FL: Academic Press, pp. 351-418.
Mann, R. 1979. The effect of the temperature on growth, physiology, and gametogenesis in manila clam Tapes philippinarum (Adams & Reeve, 1850). J. Exp. Mar. Biol. Ecol. 38:121-133.
Martoja, R. & M. Martoja-Pierson. 1967. Initiation aux techniques de l'histologie animale. Paris, France: Masson et Cie. 345 pp.
Maunder, M. N. & R. B. Deriso. 2013. A stock-recruitment model for highly fecund species based on temporal and spatial extent of spawning. Fish. Res. 146:96-101.
Meneghetti, F., V. Moschino & L. Da Ros. 2004. Gametogenic cycle and variations in oocyte size of Tapes philippinarum from the Lagoon of Venice. Aquaculture 240:473-488.
Morvan, C. & A. D. Ansell. 1988. Stereological methods applied to reproductive cycle of Tapes rhomhoides. Mar. Biol. 97:355-364.
Motavkine, P. A. & A. A. Varaksine. 1989. La reproduction chez les mollusques bivalves: role du systeme nerveux et regulation. Rapports scientifiques et techniques IFREMER 10:116-165.
Milani, L., A. Pecci, G. Ghiselli, M. Passamonti, M. Lazzari, V. Franceschini & M. G. Maurizii. 2017. Germ cell line during the seasonal sexual rest of clams: finding niches of cells for gonad renewal. Histochem. Cell Biol. 10.1007/s00418-017-1607-z.
Park, K. I. & K. S. Choi. 2004. Application of enzyme-linked immunosorent assay for studying of reproduction in the Manila clam Ruditapes philippinarum (Mollusca: Bivalvia): 1. Quantifying eggs. Aquaculture 241:667-687.
Pipe, R. K. 1987. Oogenesis in the marine mussel Mytilus edulis: an ultrastrucural study. Mar. Biol. 95:405-414.
Robert, R., G. Trut & J. L. Laborde. 1993. Growth, reproduction and gross biochemical composition of the Manila clam Ruditapes philippinarum in the Bay of Arcachon, France. Mar. Biol. 116: 291-299.
Rodriguez-Moscoso, E., J. P. Pazoz, A. Garcia & F. F. Cortes. 1992. Reproductive cycle of Manila clam, Ruditapes philippinarum (Adams & Reeve 1850) in Ria of Vigo (NW Spain). Sci. Mar. 56:61-67.
Sastry, N. A. 1979. Pelecypoda (excluding Ostreidae). In: Giese, A. C. & J. S. Pearse, editors. Reproduction of the marine invertebrates, vol. 5: pelecypods and lesser classes. New York, NY: Academic Press, Inc. pp. 113-265.
Suarez, M. P., C. Alvarez, P. Molist & F. San Juan. 2005. Particular aspects of gonadal cycle and seasonal distribution of gametogenic stages of Mytilus galloprovincialis cultured in the estuary of Vigo. J. Shellfish Res. 24:531-540.
Tang, S.-F. 1941. The breeding of the scallop [Pecten maximus (L.)] with note on the growth rate. Proc. Trans. Liverpool Biol. Soc. 54:9-28.
Uddin, M. J., K. J. Park, C. K. Kang, H. S. Kang & K. S. Choi. 2012. Annual reproductive cycle and reproductive efforts of the Manila clam Ruditapes philippinarum in Incheon Bay off the west coast of Korea using a histology-ELISA combined assay. Aquaculture 364-365:25-32.
Uddin, M. J., H. S. Yang, K. S. Choi, H.J. Kim, J. S. Hong & M. J. Cho. 2010. Seasonal changes in Perkinsus olseni infection and gametogenesis in Manila clam, Ruditapes philippinarum, from Seonjaedo Island in Incheon, off the west coast of Korea. J. World Aquacult. Soc. 41:93-101.
Valdizan, A. 2011. Bases biologiques de la proliferation d'un Gasteropode invasif de la cote Atlantique europeenne, Crepidula fornicata. These de doctorat, Nantes.
Vaschenko, M. A., I. G. Syasina, P. M. Zhadan & L. A. Medvedeva. 1997. Reproductive function state to the scallop Mizuhopecten yessoensis Jay from polluted areas of Peter the Great Bay, Sea of Japan. Hydrohiologia 352:231-240.
Weibel, E. R., G. S. Kistler & W. F. Scherle. 1966. Practical stereological methods for morphometric cytology. J. Cell Biol. 30:23-38.
Williams, M. A. 1981. Quantitative methods in biology. 2. Stereological techniques, vol 6. In: Glauert, A. M., editor. Amsterdam, The Netherlands: North Holland Publishing Company, pp. 5-84.
Xie, Q. & G. M. Burnel. 1994. A comparative study of the gametogenic cycles of the clams Tapes philippinarum (Adams & Reeve 1850) and Tapes decussatus (Linnaeus) on the south coast of Ireland. J. Shellfish Res. 13:467-472.
DAPHNE CHEREL AND PETER G. BENINGER (*)
Laboratoire de Biologie Marine, Faculte des Sciences, Universite de Nantes, 2 rue de la Houssiniere, 44322 Nantes Cedex, France
(*) Corresponding author. E-mail: firstname.lastname@example.org
TABLE 1. Phases of the Tapes philippinarum oogenic cycle. Phase Period Elements observed Gametogenesis April to August MO. IO, and AO (AMO (Fig. 4A, B) and AIO) Scarce AL and IAT Resorption (Fig. 4C) September to November Few oocytes, mostly identified as AO, AL and IAT Resting phase (Fig. 4D) December to March AL, IAT, macrophage cells No gametes Phase Other characteristics Gametogenesis Repeated declines in MO (Figs. 5-7): (Fig. 4A, B) dribble spawner Resorption (Fig. 4C) Resting phase (Fig. 4D) Inter-individual variation in the degradation of residual acini AMO = atresic mature oocytes; AIO = atresic immature oocytes; IAT = interacinal tissue; AL = acinal lumen. TABLE 2. Mean oocyte status indices during the active gametogenic period at the three study sites. Sites MAI AVF MVF Farmed 48.8 [+ or -] 5.7 33.7 [+ or -] 5.2 35.0 [+ or -] 4.4 Unfished 44.3 [+ or -] 6.4 33.2 [+ or -] 6.4 41.2 [+ or -] 8.3 Fished 41.9 [+ or -] 13.7 31.2 [+ or -] 11.2 37.5 [+ or -] 9.3 Total 45 32.7 37.9 Sites IVF N Farmed 31.4 [+ or -] 5.5 139 Unfished 25.6 [+ or -] 10.5 49 Fished 31.3 [+ or -] 10.6 44 Total 29.4 232
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|Author:||Cherel, Daphne; Beninger, Peter G.|
|Publication:||Journal of Shellfish Research|
|Date:||Dec 1, 2017|
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