Settlement and early nursery of juvenile Anadara grandis (Pelecypoda: Arcidae) under different conditions at the hatchery and ponds.
Saucedo, Pedro E.
Mazon-Suastegui, Jose M.
|Publication:||Name: Journal of Shellfish Research Publisher: National Shellfisheries Association, Inc. Audience: Academic Format: Magazine/Journal Subject: Biological sciences; Zoology and wildlife conservation Copyright: COPYRIGHT 2012 National Shellfisheries Association, Inc. ISSN: 0730-8000|
|Issue:||Date: August, 2012 Source Volume: 31 Source Issue: 3|
|Topic:||Event Code: 310 Science & research|
|Product:||Product Code: 0913030 Clams NAICS Code: 114112 Shellfish Fishing SIC Code: 0913 Shellfish|
|Geographic:||Geographic Scope: Mexico Geographic Code: 1MEX Mexico|
ABSTRACT Fifteen-day pediveliger larvae of Anadara grandis (shell
length, 230 [+ or -] 20 [micro]m (mean [+ or -] SD, n = 20) were
cultivated up to the juvenile stage by testing 2 densities: 71
larvae/[cm.sup.2] and 283 larvae/[cm.sup.2] for 37 days. During this
time, we registered morphological changes of the larvae from pediveliger
up to the juvenile stage, as well as absolute growth, and growth and
survival rates. The growth rate was 53.5 [micro]m/day at a density of 71
larvae/[cm.sup.2], with a survival of 60%; at a density of 283
larvae/[cm.sup.2], the growth rate was 33.6 [micro]m/day, with a
survival of 40%. Subsequently, the juveniles obtained were nursed for 57
days in 2 types of experimental units: Nestier trays suspended in a pond
and cylindrical upwelling containers with increasing water flow in the
laboratory. In each experimental unit we cultured 66,666 juveniles with
a shell length of 1.57 [+ or -] 0.05 mm (n = 20). In the Nestier trays,
growth was 4-6 [micro]m/day, with a survival of 15%. In the
upwelling-type units, growth was 5-6 [micro]m/day, with a survival of
93%. ANOVA revealed significant differences (P < 0.05) in juvenile
growth between the Nestier trays and the upwelling-type containers. Low
juvenile survival in Nestier trays is attributed to clogging with
wind-blown slime. However, a nursery of A. grandis juveniles in ponds
requires further research to show its economic feasibility.
KEY WORDS: mangrove cockle, Anadara grandis, juveniles, growth, survival
The mangrove cockle Anadara grandis (Broderip and Sowerby 1829) inhabits sandy bottoms adjacent to mangroves and ranges from the coasts of Baja California, Mexico, down to Peru (Keen 1971). It is subject to commercial exploitation (Mackenzie 2001), and its natural populations have had a large fluctuation because of disorganized capture and habitat destruction (Baqueiro et al. 1982, Mackenzie 2001, Felix-Pico et al. 2009). The increasing supply and demand of this species in Mexico has generated a particular interest in scientists and private producers for its study and culture, and its production has achieved precommercial levels in private laboratories in Mexico (Robles-Mungaray 2009, Guevara-Escamilla et al. 2010). However, despite the scientific and technological advances achieved in past years with respect to breeding this species, juvenile production (seeds) in the laboratory is still discontinuous and does not guarantee a sufficient and consistent supply. The major obstacles for production lie in low pediveliger larval survival during the critical development processes of settlement and metamorphosis, as well as in the high cost associated with maintaining a juvenile nursery, where a constant and abundant supply of good-quality microalgae is needed (Uriarte & Farias 1995, Mazon-Suastegui et al. 2008, Mazon-Suastegui et al. 2009). Thus, in some places of Ecuador and Mexico, A. grandis juvenile nursery maintenance and fattening has been performed in shrimp farm effluents as an alternative to reduce feeding costs associated with laboratory culture (Miranda-Baeza et al. 2006). However, there is still insufficient scientific data with respect to optimizing a species culture protocol. Therefore, in this study we assessed the feasibility of using floating sieves to fix A. grandis larvae to 2 sowing densities, as well as used 2 types of alternative systems in the juvenile nursery to advance the process of optimizing juvenile production protocols for this species.
MATERIALS AND METHODS
Setting and Early Nursery
We selected 15-day pediveliger larvae (230 [+ or -] 20 [micro]m in length), with a shell length of 192 [+ or -] 5 [micro]m length (n = 20), from an experimental culture performed in the laboratory of Centro de Investigaciones Biologicas del Noroeste (CIBNOR), La Paz, B.C.S. Mexico. During the culture, we used 6 sieves with a diameter of 67.5 cm and a nylon mesh opening of 140 [micro]m as substrate, and assessed 2 types of experimental densities: 71 larvae/[cm.sup.2] and 283 larvae/[cm.sup.2]. Both densities were managed in triplicate. The sieves were placed floating inside cylindrical-conical 5,000-L fiberglass tanks (diameter, 1.83 m; height, 1.43 m). A sprinkler system was placed in each sieve to supply a continuous mixture of 2:1:1 of the microalgae Isochrysis galbana, Chaetoceros calcitrans, Chaetoceros gracilis; an estimate was performed by cell count to have a final concentration of 60,000 cell/mL (Fig. 1). Temperature and salinity in the tanks were registered daily, and a total water exchange was performed every 7 days. During each exchange, we sampled 20 pediveliger larvae from each sieve to determine survival percentage and growth in the largest shell axis (measured in micrometers) by counting open or empty shells. At the same time, we registered the most representative morphological larval changes from pediveliger up to juvenile, including settlement and metamorphosis stages, by using a photographic camera connected to a compound microscope and a stereoscope. The experiment ended at 37 days when postlarvae or recently settled seeds reached an average size of 1.6 [+ or -] 0.23 mm in shell length (n = 20), which allowed manipulating them without fracturing their shells and starting their nursery.
[FIGURE 1 OMITTED]
Juvenile Nursery in Ponds
We placed 66,666 juveniles with a shell length of 1.57 [+ or -] 0.05 mm (n = 20) in a nylon bag of 40 [cm.sup.2] and a 700-[micro]m opening in Nestier oyster trays. We piled up 3 trays to form a culture module. Three modules were suspended in a rope crossing a rustic pond 12 x 8 x 2 m and filled to a capacity of 144,000 L with a seawater column of 1.5 m. In the same pond, juvenile yellow snapper, Lutjanus argentiventris, were cultured in floating cages. Every 3 days, 30% of the water in the pond was exchanged. Microalgae proliferation in the water was induced with agricultural urea fertilizer (0.25 mg/L) every 5 days, according to Caballero-Mandujano (2000).
Juveniles were moved to a nylon bag with a different mesh opening according to their growth. Every 7 days, the modules were washed and samples were taken at random from 40 organisms to record their percent survival. These samples were preserved in 4% formaldehyde solution to measure increase in shell length (measured in millimeters) and growth rate (measured in millimeters/day). Pond water temperature, salinity, dissolved oxygen levels, and pH were measured daily (Multisensor YSI model 86 and YSI model ph100); water transparency was also assessed with a Secchi disc every 3 days. To concentrate the microalgae from the ponds, 1-L water samples were filtered (Whatman GF/F) to identify them by direct observation with a Zeiss microscope. The number of microalgae was determined with a Coulter ZM particle counter equipped with a tube pore size of 100 [micro]m. Nursery ended at 57 culture days when juveniles reached a shell length of 4 to 6 mm.
Juvenile Nursery in the Laboratory
Maintenance of recently settled juveniles was also performed in cylindrical fiberglass containers of 5,000 L (diameter, 1.83 m; height, 1.43 m). The operation principle of these units is the use of a recirculation system and continuous seawater flow with food induced by an aerosyphon generating an upward current (Mazon-Suastegui 2005) (Fig. 2). In each container, we placed 66,666 juveniles (42 individuals/[cm.sup.2]) with a shell length of 1.57 [+ or -] 0.05 mm (n = 20) based on previous studies that show this density has been successful in early culture of the oyster Crassostrea gigas and the pectinid Argopecten ventricosus (Mazon-Suastegui 2005). Juveniles were fed daily with a mixture of 2:1:1 of the microalgae I. galbana, C. calcitrans, and C. gracilis at a concentration of 60,000 cells/mL. Temperature was regulated at the same value registered in the pond, using submersible heaters of 250 W. At the same time, we registered salinity, dissolved oxygen levels, and water pH data in the containers. We took samples of 40 organisms at random, for which we registered percent survival, and preserved them in 4% formaldehyde solution to measure absolute growth in shell length and growth rate. As in the pond, nursery in the laboratory ended at 57 days when the organisms reached sizes between 4 mm and 6 mm in shell length.
[FIGURE 2 OMITTED]
With shell length data we determined growth through a regression analysis. Subsequently, a 1-way ANOVA was used followed by Tukey's median comparison trial a posteriori when necessary to determine existence of significant differences in organism growth among treatments. We also compared organism growth among replicates of the oysters cultured in trays and in upwelling units. For the trial, significance was set a P [less than or equal to] 0.05. Data are presented as mean [+ or -] SD (n = 20).
Setting and Early Nursery
Growth of A. grandis pediveliger up to juvenile in the 2 experimental densities (Fig. 3) was significantly different (P < 0.05), and was adjusted to a significant linear model (r > 0.98). Growth rate was 53.5 [micro]m/day at the density of 71 larvae/[cm.sup.2], and 33.6 [micro]m/day at the density of 283 larvae/[cm.sup.2], with a survival of 60% and 40%, respectively. Development time and morphological changes of A. grandis pediveliger larvae up to juvenile stage are shown in Table 1 and Figure 4.
At the start of the culture, pediveliger larvae with an eyespot, a shell length of 230 [+ or -] 20 [micro]m (n = 20), and a shell width of 192 [+ or -] 5 [micro]m showed concave transparent valves, a robust umbo, and an active velum. The eyespot was black and ovoid, and 22 [+ or -] 5 [micro]m in its longest diameter (n = 20; Fig. 4A). During metamorphosis, larvae showed natal and inactive behavior. The foot developed and the velum lost the cilia gradually. Metamorphosis originated in benthic postlarvae (length, 320 [+ or -] 20 [micro]m; width, 270 [+ or -] 10 [micro]m) that adhered and crawled around the substrate with the foot without the velum. Dissoconch secretion started at the extreme lateral parts of the valves (Fig. 4B). At a length of 350 [+ or -] 10 [micro]m and a width of 270 [+ or -] 10 [micro]m, secretion of the first ornaments started in the form of extended growth processes in the prodissoconch 21 [+ or -] 5 [micro]m in length (Fig. 4C). The dissoconch surrounded the prodissoconch periphery, and when the dissoconch secretion increased, growth lines appeared. In postlarvae 490 [+ or -] 10 [micro]m in length and 390 [+ or -] 10 [micro]m in height, ornaments in the dissoconch measured from 72-142 [micro]m in length. Secretion in the dissoconch was greater in the extreme lateral sides of the valves as the shell acquired an extended form. Margin rib development could be observed in the shell (Fig. 4D). At this size, some individuals still showed the eyespot. Ribs developed from the umbo up to the distal margin of the shell in juveniles 958 [+ or -] 10 [micro]m in length. The contracted siphons could be observed as 2 dark spots through the shell. The shell border was crenulated (Fig. 4E). At a size of 1,707 [+ or -] 10 [micro]m in length and 1,075 [+ or -] 10 [micro]m in height, 26 ribs formed and were separated by deep interspaces (Fig. 4F). At the level of the umbo, central, prominent, and elevated convex growth started in the shell. Ornaments all over the shell formed a dark-brown periostracum. At 37 days, juveniles 1,830 [+ or -] 23 [micro]m in length and 1,260 [+ or -] 17 [micro]m in height had a square shell with a central elevation (Fig. 4G), and they showed adult morphological characteristics. At this culture size and time, larval settlement up to juvenile ended. Shell hardness allowed manipulation of juveniles to start the next nursery culture stage.
[FIGURE 3 OMITTED]
Juvenile Nursery in Ponds
Juvenile growth during nursery in Nestler trays in ponds is shown in Figure 5. The regression analysis showed that growth adjusts to a significant logarithmic model (r [greater than or equal to] 0.94). In each module, we harvested an average of 30,800 juveniles, with a size of 2.1-4.6 mm. In the upper tray of the experimental models, the highest growth rate occurred (58 [micro]m/day) at a size range of 2.9-4.61 mm in shell length and a survival rate of 19%. In the intermediate trays, the growth rate was 50 [micro]m/day with shell length range of 2.9-4.5 mm and a survival rate of 16%. In the lower trays, growth rate was 42 [micro]m/day with a shell length of 2.1-3.3 mm and a survival of 11%. These differences were significant among the different levels of the Nestier trays (P < 0.05). Tukey's multiple range test showed that growth was different among trays in the module.
From the total particles and organisms suspended in the pond water, 60% corresponded to the microalga Nannochloropsis sp. (85,000 [+ or -] 15,000 cells/mL); 32% to fine slime; and 8% to copepods, rotifers, and ascidia, in the water column, average transparency was 0.50 m; water temperature, 24 [+ or -] 3[degrees]C; salinity, 37-39 ppt; dissolved oxygen, 6.9 [+ or -] 0.3 mg/L; and pH, 7.6 [+ or -] 0.2.
Juvenile Nursery in the Laboratory
Growth of A. grandis juveniles in the upwelling unit showed a very different trend from the pond (Fig. 6), although it was also adjusted to a significant logarithmic model (r [greater than or equal to] 0.95). In upwelling unit 1, we recorded a growth rate of 50 [micro]m/day, with a final average size of 4.39 [+ or -] 0.63 mm and a survival rate of 92%. In upwelling unit 2, we recorded a growth rate of 60 [micro]m/day, an average final size of 4.9 [+ or -] 059 mm, and a survival rate of 95%. Everything was lost in upwelling unit 3 because of an electrical problem with the heaters. In the upwelling units, water temperature was 24 [+ or -] 1[degrees]C; salinity, 37-39 ppt; dissolved oxygen, 6.6 [+ or -] 0.4 mg/L; and pH, 7.8 [+ or -] 0.1. Tukey's multiple range analysis showed there were no significant differences in juvenile growth among treatments in upper Nestier tray modules and upwelling unit 1. Juvenile growth in lower Nestler tray modules had a significant difference among all nursery treatments.
[FIGURE 4 OMITTED]
In the majority of bivalve mollusc species, the plantigrade postlarval stage is a transition between the swimming planktonic life of veliger larvae with an eyespot and the sedentary benthic life of juveniles (Bricelj & Shumway 1991). A high natural mortality occurred in this transition as a result of the lack of sufficient energy reserves to complete metamorphosis (Loosanoff & Davis 1963). In the current study, the pediveliger stage of A. grandis was present in postlarvae with a shell length of 230 [+ or -] 20 [micro]m at 15 days after the veliger stage and 7 days after the eyespot appeared (Table 1). This transition time was similar to that reported in Anadara tuberculosa postlarvae that were 233 [micro]m in shell length (Reynoso-Granados et al. 1999) and Anadara granosa postlarvae that were 240 [+ or -] 10 [micro]m in shell length (Wong & Lim 1985). In species of the genus Anadara, metamorphic transition times can be considered extended because, in other bivalves like pectinids, transition occurs between 24 h and 72 h (Bricelj & Shumway 1991). In fact, in the majority of the bivalve species, plantigrade postlarvae can differ from swimming veliger larvae in the secretion of the dissoconch. In this study, dissoconch characteristics of A. grandis from the shell form and superficial growth processes (ornaments) to the growth lines were identified starting from the formation of the 26 ribs and the periostracum, as well as by the formation of a square shell with central elevation. These are the typical characteristics of the species that distinguish it from other Anadara species that are cultured in Latin America (Brusca & Brusca 2005).
[FIGURE 5 OMITTED]
[FIGURE 6 OMITTED]
Currently, in aquaculture laboratories in the Republic of El Salvador, A. tuberculosa plantigrade postlarvae have a survival rate of 15% in culture conditions (Vasquez et al. 2009) using artificial nylon collectors that have been efficient in fixing pectinids (Narvarte et al. 2002). In other experimental cultures of A. tuberculosa, postlarval survival has been null in oyster trays at 14 culture days (Reynoso-Granados et al. 1999). However, the results of our work show that the use of floating sieves, similar in form to oyster trays, was efficient as a fixing substrate for A. grandis as a survival of 40% and 60% was obtained in the 2 densities assessed, respectively. This survival percentage is high with respect to the 15% reported for A. tuberculosa (Vasquez et al. 2009), 10-30% registered in pectinids (Nicolas et al. 1995), and 30-35% reported in Crassostrea iridescens cultured in trays (Robles-Mungaray 2009).
During the A. grandis settlement process, experimental densities of 71 larvae/[cm.sup.2] and 283 larvae/[cm.sup.2] could seem high when compared with those used for the Catarina clam A. ventricosus (Blacio et al. 2001) and C. iridescens (Robles-Mungaray 2009), which were 16-20 larvae/[cm.sup.2] and 6-8 larvae/[cm.sup.2], respectively. For A. grandis, we calculated an approximate density of 4 and 15 times higher than that for A. ventricosus as the low and high density, respectively. In fact, A. grandis survival was high in the sieves with high density, which could be an indication that these larvae could be cultured intensively with good results, based on previous studies in which it was reported that densities of 16-20 larvae/[cm.sup.2] have been used in A. grandis hatcheries (Vasquez et al. 2009).
A determining difference factor in survival percentages of other mollusc postlarvae was the competition for space, which resulted in major fixation at a low density and a lesser fixation at a high density (Brand et al. 1980). In consequence, postlarval growth might decrease when density exceeds a certain value of a determined size for A. grandis fixation. Different works on bivalve culture in nurseries have shown that the growth rate is related to environmental factors, as seen in A. granosa in a tropical environment (Broom 1983, Broom 1985). In our study, temperature (23-25[degrees]C), salinity (37-39 ppt), dissolved oxygen (6.6-7.2 mg/L), and pH (7.4-7.8) levels are similar to those reported by Miranda-Baeza et al. (2006) for A. grandis at a shell length of 50 [+ or -] 0.37 mm, where the highest filtration and clarification rates were obtained at 25[degrees]C, with pH and oxygen levels of 7.9 [+ or -] 0.2 and 6.8 [+ or -] 0.4 mg/L, respectively. In this study, variation ranges in environmental factors were similar in 2 nursery systems. They corresponded to the habitat where the species of the genus Anadara are naturally distributed within the lagoon systems of Mexico, where the climate is hot and humid, and the surrounding vegetation is typical of mangrove systems (Antoli & Garcia-Cubas 1983).
In bivalve nursery systems, another factor related to juvenile growth is food availability (Uriarte & Farias 1995). In the current study, the microalgae Nannochloropsis sp. was abundant and constant as a result of the fertilization performed in the polyculture pond with marine fish. In the 2 nursery systems, growth rates were 53.3 [+ or -] 6.0 [micro]m/day and 63.3 [+ or -] 10.0 [micro]m/day, indicating that A. grandis has an intermediate growth in relation to other species of the genus Anadara, such as Anadara broughtonii, which has the fastest growth rate (110 [micro]m/day) (Yoo 1977, Kim et al. 1982), whereas others have lesser rates, such as A. granosa (33 [micro]m/ day (Broom 1985)) and Anadara seniles (26.6 [micro]m/day (Debenay et al. 1994)). Only Anadara subcrenata showed an intermediate growth from 56.666.6 [micro]m/day (Ting et al. 1972), similar to that observed in the current study.
In the laboratories of the Republic of El Salvador, a maximum survival rate of 90% has been obtained in nursery experiments of A. tuberculosa (Vasquez et al. 2009). Based on this information, a survival of 85% can be considered acceptable during nursery in upwelling cylinders. However, a nursery survival of 11-19% in Nestier trays in the pond with marine fish was low, which could be attributed to slime accumulation suspended in the water column of the pond and carried by the wind, which clogged the trays when sediments settled, particularly the lower ones (pers. obs.). In the higher trays of the module associated with less slime accumulation, we obtained the highest survival rate of A. grandis. Similar results were obtained in a fattening experiment of A. tuberculosa in oyster trays suspended in a platform over a mangrove channel (Villalobos & Baez 1983). According to Villalobos and Baez (1983), survival and growth were higher in juveniles cultured in the platform without clogging than in juveniles immersed in mud all the time. In pectinid cultures in trays suspended in ponds, water flow in the pond and within the culture trays has been an important factor in organism response, which is a result of the negative effect of water flow reduction and, in consequence, food availability inside the culture artifacts. These are factors that need to be considered in future cultures of A. grandis in ponds, as it has been demonstrated in a nursing research study of the pectinid A. ventricosus in the sea using Nestier trays (Maeda-Martinez et al. 1997). Based on these considerations and according to the results of our work, we suggest that nursery of A. grandis juveniles be performed in ponds in upwelling units to promote an acceptable growth of the species.
We thank C. Aldana-Aviles and M. Vargas-Mendieta for microalga culture, D. Dorantes-Salas for English translation and editing, and G. R. Hernandez-Garcia for managing the Corel images.
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TEODORO REYNOSO-GRANADOS, (1) * PABLO MONSALVO-SPENCER, (1) PEDRO E. SAUCEDO, (1) JOSE M. MAZON-SUASTEGUI (1) AND MIGUEL ROBLES-MUNGARAY (2)
(1) Programa de Acuicultura, Centro de Investigaciones Biologicas del Noroeste, S.C. Mar Bermejo 195, Col. Playa Palo de Santa Rita, La Paz, B.C.S., Mexico, (2) Acuicultura Robles, S.P.R., Privada Quintana Roo no. 4120, La Paz, B.C.S., Mexico
* Corresponding author. E-mail: email@example.com
TABLE 1. Time of development stages of Anadara grandis from pediveliger larva to juvenile under laboratory conditions at 24-25[degrees]C. Time 0 = Culture start; d = days. Culture Time (days) Stage Figure 4 0 Pediveliger larva with eyespot A Planktonic stage with start of settling stage Rudimentary extensible foot Active velum 2 Metamorphosis Foot in development Loss of velum cilia 7 Post larva B Benthic stage Functional foot Without velum Secretion of dissoconch 9 First ornaments (thorn shape) C in prodissoconch Formation of growth lines 15 Formation of ribs in shell margin D 19 Increase in number of ribs E Crenulated shell border 27 Juvenile F Ribs formed (n = 26), separated by deep interspaces Elevated convex growth under the umbo in the shell center Ornaments form a dark-brown tone in the periostracum 37 Square shell with central elevation, G morphological characteristics of an adult of the species Time 0, culture start.
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