Printer Friendly

Larval quality of a nonnative bivalve species (European oyster, Ostrea edulis) off the east Canadian coast.

ABSTRACT A population of European oysters (Ostrea edulis) was recently discovered off the eastern coast of Canada. The occurrence of this newly established population led various aquaculture stakeholders to consider this species commercially. To better assess recruitment capacity, the physiological quality of the adults, eggs, and larvae was described using glycogen and various lipid contents, including a TAG/ST ratio. Adults, early nonfeeding stages (eggs, pretrochophores, trochophores, veligers) and newly-released larvae were periodically sampled in the wild and/or the hatchery during the spawning period. The initial glycogen content was higher in wild oysters than hatchery-conditioned ones. The following larval stages showed higher lipid content in wild individuals as well. Spat collection in the wild was abundant. The free-living larvae, however, did not survive more than 10 days in the hatchery. TAG/ST ratios decreased during the veliger stage development and were lower when larvae were released from the paleal cavity of the female. The high spat collection observed in the wild suggests that observed TAG/ST ratios in early nonfeeding larvae are representative of a good larval quality. The physiological quality of the larvae in the wild seems to be good enough to allow larvae to settle in this particular environment.

KEY WORDS: nonnative species, Ostrea edulis, European oyster, conditioning, early nonfeeding stages, lipid analysis


Several nonnative species of oysters were introduced in various countries during the past decades (e.g., Crassostrea angulata in Spain, Andrews 1980), Crassostrea gigas in France (Grizel & Heral 1991), Ostrea edulis in the United States (Welch 1964, Medcof 1961). Most of these introductions attempted to fill an economic loss associated with the decline of commercially important indigenous species. Several introduced species became successfully established in their new habitat, whereas others failed. Failed introductions could be attributed to nonfavorable environmental conditions and/or habitat quality (Eldredge 1994).

The establishment of a nonnative species in a new habitat essentially depends on the success of recruitment processes. These processes are highly variable from year to year and are related to many factors (Brand et al. 1980). These factors include larval behavior (Wehrtmann 1990, Miron et al. 2000), predation (Andre et al. 1993, Dame 1996) and hydrodynamics as well as various physical, biological and chemical factors that occurred in the benthic habitat (Abbe 1986, Navarrete & Castilla 1990). Larval settlement and recruitment are, for instance, strongly related to the physiological quality of the larvae (Gallager et al. 1986, Millican & Helm 1994, Miron et al. 2000, Pernet et al. 2005). Larval quality refers to the physiological condition and the capacity of a larva to grow and survive under various environmental conditions during its life cycle (Racotta et al. 2003).

All stages of larval development are energy consuming processes (Helm et al. 1973, Bayne 1976, Holland 1978, Gallager et al. 1986, Gallager & Mann 1986, Rodriguez et al. 1990). A larva, for instance, will be particularly susceptible to life-threatening energy deficits during metamorphosis (Haws et al. 1993). Energy is stored during 2 stages of development: (1) during the embryonic development, when genitors provide endogenous reserves to the eggs (Bayne 1973, Helm et al. 1973, Walne 1974, Bayne et al. 1978, Lannan 1980, Lannan et al. 1980, Gallager et al. 1986, Gallager & Mann 1986) and (2) during feeding when newly-liberated larvae rely on phytoplankton until settlement and metamorphosis (Whyte et al. 1989, 1990). Several studies showed that lipids are the most metabolized constituents during metamorphosis. Lipids are accumulated in preference compared with proteins and carbohydrates (Holland & Spencer 1973, Bayne et al. 1975, Lucas et al. 1986).

Although the use of proteins has been documented (Bartlett 1979, Rodriguez et al. 1990), several researchers suggest that lipids represent the major energy reserve during larval development (Millar & Scott 1967, Helm et al. 1973, Holland & Spencer 1973, Holland & Hannant 1976). Neutral lipids are used primarily when food is scarce as well as during metamorphosis (Holland & Spencer 1973). Helm et al. (1973) found that the initial growth rate of larvae may be positively correlated with the lipid content of the eggs and that larval development and settlement success is positively related to lipid levels.

The recent discovery of a European oyster population in New Brunswick (Canada) waters led various stakeholders to consider this species commercially. This population reproduces naturally and may potentially produce seed for aquaculture purposes whether environmental conditions allowed it. Our knowledge of its reproductive potential, settlement behavior, and recruitment is still limited in this particular environment. This population is occurring at the boreal limit of its distribution on the western side of the Atlantic where environmental conditions are extreme and variable during a given year, as well as through the years. It is possible that environmental conditions are not suitable enough to allow an adequate spat collection for culture operations.

The aim of this study is to evaluate the physiological quality of adults by glycogen content and various larval stages by lipid class levels to better estimate the spat collection potential of the European oysters off the coast of New Brunswick. A triacyl-glycerol/sterol (TAG/ST) ratio will be used to evaluate the larval quality of early-non feeding stages produced from wild oysters. This particular ratio has already been used to evaluate the quality of bivalve, crustacean, and fish larvae under different nutritional and pollutant stresses (Fraser 1989, Harding & Fraser 1999, Pernet et al. 2003). Because of the lack of data on early non- feeding stages for this species, we also reared oysters larvae produced from the same population in the hatchery. These data will be used as a comparison tool to describe the physiological quality of larvae.


Treatment of Broodstock and Spawning

Oysters (mean size of 101.5 mm x 95.4 mm) were collected on April 28, 2005. Individuals were dredged from Lockhart Lake (45.67735[degrees]N, 64.74505[degrees]W) near New Horton (NB), Canada. This water system (salinity: 25.4 [per thousand] to 29.8 [per thousand], depth: 1-12 m) covers an area of approximately 69 hectares and is influenced by the tide cycle of the Bay of Fundy (Bataller et al. 2006). The bottom is primarily composed of mud. A total of 410 oysters were randomly allotted to 10 Vexar bags (41 individuals/bag) of 85 x 9 x 44 cm. Five bags were sunk to the bottom of the lake at two experimental sites (Sites A and B, Fig. 1). A total of 205 oysters were collected at the same time and transported to the Bedford Institute of Oceanography (BIO) in Dartmouth (NS). These oysters were cleaned and randomly allotted to five Vexar bags (41 individuals/bag) upon arrival. The bags were then placed in a conditioning tank (96 x 76 x 85 cm). The conditioning of oysters began at 7[degrees]C on 29 April 2005. The seawater temperature was raised gradually until it reached 18[degrees]C during a period of 35 days. The seawater was pumped from Bedford Basin at a depth of 20 m and passed through a sand filter to remove particles >20 [micro]m. The brood-stock was fed on a mixed ration of algae at similar proportion of the following species: Isochrysis sp., Tetraselmis sp., Pavlova lutheri, Thalassosiera weissflogii, Chaetoceros muelleri, Skeletonema costatum, and Tetraselmis striata and maintained daily at a density of 40-50 cells [micro][L.sup.-1]. Algae were added continuously into the seawater by an electromagnetic dosing pump. The adult oysters were conditioned under constant conditions of salinity (30 [+ or -] 1 [per thousand]), dissolved oxygen concentration (95%) and pH (8).

Animal Sampling

Adult Oysters

Five oysters (one per bag) were sampled every two weeks from June 9 to September 21 at each site in Lockhart Lake (Fig. 1). Five oysters were sampled similarly in the hatchery at the same time. The flesh of these oysters was frozen at -20[degrees]C until glycogen analysis.

Early Nonfeeding Stages

The number of oysters sampled was raised when broods of oysters were observed in the wild and in the hatchery. Thirty individuals were then sampled at three-day intervals during the spawning period to maximize the number of eggs and larvae for lipid analysis. Three different stages were sampled: (1) pretrochophores, (2) trochophores (individuals from early shell-plate development to ones with shell valves that completely enclosed the body), and (3) completely developed veligers (Helm et al. 1991). Once a brooding oyster was found, 20 thousands eggs or larvae were collected in triplicate according to Utting & Spencer's (1991) protocol and then filtered onto a prebaked (450[degrees]C) GF/C filter. These filters were stored in 1-mL dichloromethane in amber glass vials with Teflon liner caps under a nitrogen atmosphere at -80[degrees]C until lipid extraction. The remaining eggs or larvae were preserved in a 70% alcohol solution to measure their surface area ([micro][m.sup.2]).

Newly-Liberated Larvae

This larval stage was sampled only in hatchery conditions. When newly-liberated larvae were released from the mother's paleal cavity, individuals were flowed through a sieve (125 [micro]m) positioned under the seawater outflow pipe of the tank. Twenty thousands larvae were collected from each larval release and preserved following the method described earlier. The remaining larvae were transferred (3 larvae [mL.sup.-1]) in a conical rearing tank (2001) filled with seawater maintained at 21 [+ or -] 1[degrees]C. The tank was drained and refilled every two days. The D-larvae were fed daily with a 1:1 standard mixture of the flagellates Isochrysis sp. (clone T-ISO, termed Tahitian Isochrysis) and Pavlova lutheri (clone MONO) at a density of 30-35 cells [micro][L.sup.-1]. Six larval releases occurred in the hatchery, but only three of the six survived more than 10 days in the rearing tank. After 8 days of growth, 10,000 larvae were collected and preserved from each larval release. Newly-liberated larvae from the wild were not sampled because of potential contamination from other larvae (bivalves and/or gastropods), diatoms and other debris of similar size, which will give erroneous lipid content data.

Laboratory Analyses

Glycogen Analysis

The flesh (6.73 g [+ or -] 2.46 g) of the frozen oysters was grounded and homogenized in a sodium citrate solution (1/10 w/v, 100 mM). A volume of 1.5 mL was thereafter taken from each sample and transferred to a clean tube, which was placed on a YSI 2710 Turntable coupled with a YSI 2700 Select Biochemistry Analyzer (YSI Inc, Yellow Springs, OH). Three 25 [micro]L replicate samples were analyzed for glycogen content (g [L.sup.-1]). A volume of 5 mL of the homogenate was dried at 100[degrees]C for 12 h to determine the dry weight of flesh. This additional data were used to determine the glycogen content as mg glycogen [(g of dry weight of flesh).sup.-1].

Lipid Extraction

Lipids were extracted from eggs and larvae after a maximum of six months storage period. Eggs and larvae samples were grounded three times in 1.5 mL of C[H.sub.2][Cl.sub.2]-MeOH (2:1 v/v) in an ice bath to remove the organisms from the filter. KCl (0.88%) was added to the previous solution to obtain C[H.sub.2]-[Cl.sub.2]-MeOH-KCl (2:1:0.6; v/v/v; Folch et al. 1957). The homogenates were mixed and centrifuged at 4,000 rpm for 2 min to obtain a biphasic system. The lipid fraction (lower phase) was removed and transferred to a clean tube. The solvent was evaporated under a nitrogen flow and lipids were thereafter suspended in 0.1 mL C[H.sub.2][Cl.sub.2].


Lipid Class Composition

Lipids (2% to 5% of total extraction depending on eggs or larvae sample) were spotted onto the S-III Chromarods (Iatron Laboratories Inc., Tokyo, Japan) using a Hamilton syringe. Four different solvent systems were used to obtain three chromatograms per rod (Parrish 1987). This method allows the separation of various lipid compounds including triacylglycerols (TAGs), free sterols (STs), and phospholipids (PLs). The total lipid (TL) content was also considered and represented the sum of all lipid classes mentioned earlier and the following ones: aliphatic hydrocarbons (HCs), free fatty acids (FFAs), ketones (KETs), free fatty alcohol (ALCs), diglycerides (DGs) and acetone mobile polar lipids (AMPLs). Chromarods were scanned by flame ionization detection with a Iatroscan Mark-VI analyzer (Iatron Laboratories Inc., Tokyo, Japan). Lipid classes were identified and quantified by the use of standard calibration curves obtained for each lipid class. Within each set of rods, one was used for the lipid standard and another one for the extraction blank.

TAG/ST ratio was used as a larval quality marker. Analyses were performed on 42 broods of larvae. Twenty-four broods were obtained from Lockhart Lake. Ten were assessed as pretrochophores, seven broods were classified as trochophores and seven were finally assessed as veligers. The remaining 18 broods were obtained from the hatchery. Five were assessed as pretrochophores, six as trochophores and seven as veligers. Six replicate samples of newly-liberated larvae and three replicate samples of 8 dpl (day postliberation) collected from the hatchery were also analyzed for lipid content.

Growth Measurements

The area of 100 eggs or larvae from brooding females (wild: pretrochophores from 5 females, trochophores from 3 females, veligers from 5 females; hatchery: pretrochophores from 3 females, trochophores from 3 females, veligers from 2 females) were measured using a compound microscope (BX41TF, Olympus, Tokyo, Japan) at a magnification of x 10 with an image capture camera (Evolution VFfast, Media cybernetics, Silver Spring, Canada) and Image processing software (Image-Pro plus V5.0, Media cybernetics, Silver Spring, Canada).

Data Analysis

A linear regression was used to evaluate the glycogen content in wild and hatchery-conditioned oysters. The difference between wild and hatchery (wild-hatchery) glycogen content was used as the dependent variable. The independent variable was expressed as Julian days. Outliers were examined using Cook's distance (Cook 1977).

Two one-way MANOVAs were used to investigate the lipid class composition (TAG, ST, PL, and TL) of the eggs and larvae. These distinct MANOVAs were used to evaluate field and hatchery conditions independently because of obvious conditioning differences and also because it was an experiment without replication. These analyses also helped evaluate the lipid class composition between each female for the various studied stages. A logarithm transformation was performed on TAG, whereas a square root transformation was performed on ST, PL, and TL. Where differences were detected, Tukey-Kramer multiple comparison tests were used to determine which means was significantly different. A one-way ANOVA was performed on rank-transformed data to evaluate the surface area of wild and hatchery eggs and larvae.

Homoscedasticity was tested using Levene's test and confirmed by graphical examination of the residuals. A significant threshold of 0.05 was adopted for all statistical tests. All statistical analyses were run using SAS (SAS Institute 1982). All data in figures are presented in their original, untransformed scale.



Gametes differentiation occurred in mid June (12[degrees]C) in wild oysters. Females brooding larvae were first observed in the wild on July 14 (18[degrees]C) and remained present until August 23 (17[degrees]C). This suggests that spawning lasted about 6 wk. Similar results were observed in the hatchery, whereas brooding females were first observed on July 5 and remained present until August 12. Water temperature, however, reached 18[degrees]C on 3 June in the laboratory.

Glycogen concentrations ranged from 1.22-50.45 mg glycogen [(g dry weight of flesh).sup.-1] in wild oysters, and from 3.59-19.82 mg glycogen [(g dry weight of flesh).sup.-1] in the hatchery. As expected, glycogen content was higher in oysters sampled early in June than later during the summer (Fig. 2). There was, however, an important difference in the initial glycogen content in wild [(33.22 mg glycogen [(g dry weight of flesh).sup.-1]] and hatchery-conditioned oysters [(17.62 mg glycogen [(g dry weight of flesh).sup.-1]]. The linear regression showed that differences in glycogen content were stronger in early June. These differences decreased through time. A rapid decrease in glycogen content (wild: 66.38%; hatchery: 51.46%) was observed between June and July corresponding to the gametogenesis of the oysters. The glycogen contents observed in wild and hatchery-conditioned oysters were almost similar immediately after this rapid decrease (on July 6). No significant glycogen recovery was noted during the spawning period. The lower glycogen content observed in the oyster tissue was observed at the end of July [8.87 and 6.91 mg [(g dry weight of flesh).sup.-1] in wild and in hatchery, respectively]. The glycogen content of hatchery-conditioned oysters remained constant from July until the end of August with no significant variations. However, the glycogen content difference between wild and hatchery-conditioned oysters tends to reverse at the beginning of September.


Lipid Composition of Larvae


The one-way MANOVA outlined a significant difference (P < 0.001) in the lipid content of early nonfeeding wild larvae. Tukey-Kramer test showed that PL content between pretrochophore and veliger stages was significantly different (P = 0.007) with higher values occurring in the pretrochophores (Table 1, Fig. 3). The content of the other lipid classes such as TAG and ST did not differ significantly (P = 0.236 and P = 0.726, respectively). Though the MANOVA failed to find a significant difference, our results still showed that TL content tend to be different during the growth of early nonfeeding larvae. TL content went from 109.43 ng [larva.sup.-1] in pretrochophores to 101.77 ng [larva.sup.-1] in trochophores and to 83.79 ng [larva.sup.-1] in veligers.


The MANOVA showed that the lipid content of the early nonfeeding and newly-released hatchery larvae was significantly different (P < 0.001). TAG (P < 0.001) and TL (P = 0.021) contents varied in eggs and larvae, whereas ST (P = 0.199) and PL (P = 0.092) contents did not (Table 2, Fig. 3). In general, TAG content in early nonfeeding stages was significantly different (pretrochophores: P < 0.001; trochophores: P < 0.001; and veligers: P < 0.001) from the one observed in newly-released larvae. An important decrease in lipid content was observed during the transition from the veliger stage to newly-liberated larvae. About 82% of the TAG was metabolized during this period. The ST and PL lipid classes showed a decrease of 27.30% and 40.26%, respectively. TL content also decreased from 60.25-27.55 ng lipids [larva.sup.-1]. A nonsignificant increase in lipid content was thereafter observed in 8 dpl larvae.

Hatchery Versus Wild

The comparison between the lipid content of eggs and larvae from wild and hatchery-conditioned oysters could not be statistically demonstrated. However, our results showed that important differences in larval lipid contents may be observed. The energetic reserves (TAG) and structural lipids (PL and ST) of wild larvae are two times higher than those from the hatchery. The TL content in pretrochophores, for instance, was 109.43 ng [larva.sup.-1] in wild oysters and 51.21 ng [larva.sup.-1] in hatchery ones (Fig. 3). The lipid class contents followed the same tendency in trochophores and veligers. Our results also suggest that lipid class contents tend to decrease during the early larval development in the wild whereas it tends to increase in the hatchery. A one-way ANOVA performed on rank transformed data (Table 3) showed that the area ([micro][m.sup.2]) of the eggs and larvae was also significantly different depending on their culture environment. This was observed for the pretrochophores (P < 0.001) and veligers (P < 0.001), but not in the trochophores (P = 0.112). The absence of a significant difference in the trochophores is probably caused by a higher variability of the area in the hatchery. The area of the pretrochophores, for instance, was about 10,959 [micro][m.sup.2] and 9,110 [micro][m.sup.2] for wild and hatchery-conditioned oysters, respectively. The area difference in other stages was, however, less apparent.

Larval Quality

TAG/ST ratios varied throughout the larval development (Fig. 4). An initial TAG/ST ratio of 12.49 was observed in wild oysters for the pretrochophores. It then decreased with subsequent stages passing from 10.62 in the trochophores to 7.72 in veligers. The same tendency was observed in the hatchery oysters. An important decrease in TAG/ST ratios was observed during the transition from the veliger stage to newly-liberated larvae. A small increase in TAG/ST ratio was thereafter observed once the larvae were released in the water column until they aged 8 days, coinciding with the time that free-living larvae began to feed on exogenous sources.



The life cycle of marine bivalves is strongly related to environmental parameters such as temperature and food availability as well as to cycles of storage and utilization of biochemical substrates (Taylor & Venn 1979, Zandee et al. 1980, Beninger & Lucas 1984, Arellano-Martinez et al. 2004). In general, when food is abundant, energetic reserves accumulate prior to gametogenesis and are subsequently used in gamete production (Mathieu & Lubet 1993). Glycogen is known to be a major energetic reserve for adult bivalves (Bayne & Newell 1983, Gabbott 1983, Gade 1983, Akberali & Trueman 1985, Frolov & Pankov 1992a). In our study, the initial glycogen content of wild European oysters was higher than the ones kept in the hatchery. Beukema et al. (2001) reported that an individual with high glycogen content may lose more weight during gametogenesis, without compromising growth and survival, than an animal with a low glycogen content. Our results tend to support this observation. Wild oysters showed a higher loss of glycogen during gametogenesis than hatchery ones. This difference further suggests that wild oysters had a higher reproductive effort and higher egg quality than hatchery one. This was later confirmed by important differences in lipid contents observed in pretrochophore stages. Such an initial glycogen content difference between wild and hatchery-conditioned oysters could be explained by the conditioning method, which may affect the timing of gametogenesis and spawning processes. The success of these events depends on the physiological condition and particularly the prespawning condition of the adults (Hendriks et al. 2003).


Gametogenesis and spawning are affected by temperature (Korringa 1947, Sato 1951, Marteil 1960, Le Dantec 1968, Mann 1979). In the present study, gametogenesis was observed at 12[degrees]C. Brooding oysters were observed at 18[degrees]C. These events occurred within a 35-day interval in the wild. In the hatchery, these temperatures were observed 13 and 32 days after the initiation of conditioning, respectively; this suggests that wild oysters feed during a longer period of time compared with those kept in the hatchery and consequently accumulated more biochemical substrates.

The pretrochophore stage is known to last 3-3.5 days, the trochophore stage 1.5-2 days or less (Orton 1936). The veliger stage, which is more variable, may last 4 days or less; this suggests that the first pretrochophores appeared in the hatchery between the 23 and 27 days after 18[degrees]C was reached. The delay observed in the spawning period in the hatchery could be explained by the fact that oysters were not yet ready to spawn at 18[degrees]C. The temperature might have increased too quickly, which in turn might have stressed the oysters. Gallager & Mann (1986) reported that a life cycle interruption while using artificial conditioning at a high temperature, could force the production of a lower number of eggs and/or the production of eggs of suboptimal quality. Martinez & Perez (2003) observed, in a hatchery, a gonadosomatic index decrease in Argopecten purpuratus when the increase in temperature ceased, resulting in a gonad maturation delay. An increase in temperature during the previtellogenic period may produce an increase in metabolism, stimulating the use of energy reserves otherwise required for subsequent vitellogenesis (Griffond et al. 1992).

Lipid Composition of Larvae

Early Non-feeding Stages

Lipids are considered as a major energy resource to sustain the embryonic and larval development of many marine bivalve species (Holland & Spencer 1973, Holland 1978, Fraser 1989). Our results showed that there was no important variation in the content of various lipid classes (e.g., TAG, ST) as well as in TL during the early nonfeeding larvae development. Only the PL content differed significantly between wild pretrochophores and veligers. Several studies reported that PL could play an energetic role during the early larval development or starvation period (Tocher et al. 1985, Fraser et al. 1988, Fraser 1989, Helm et al. 1991, Delauney et al. 1992).

Although the TL contents observed in the early nonfeeding stages were not significantly different, we observed a 23% TL reduction between pretrochophores and veligers. This decrease is comparable to results obtained by other researchers on various species. Waldock (1979), in Helm et al. (1991), found a decrease of 19%, within a 24-h period, in TL content between fertilized eggs and shelled D-larva in Crassostrea gigas. Gallager et al. (1986) reported a lipid loss of 70% of parentally derived TL, principally TAG, during the development of the eggs of Mercenaria mercenaria and Crassostrea virginica within the same time period. The lipid utilization observed in this study was different from those observed in the hatchery ones, because the TL content increased by 15% during the early nonfeeding larvae development. This increase did not differ significantly and could be explained by the variability of lipid content from samples collected in the hatchery. Helm et al. (1991) observed a TL content of 71 [+ or -] 7 ng [egg.sup.-1] in an oyster population from Poole Harbour, UK. The TL contents observed in their hatchery oysters were similar. Our hatchery results showed that TL content found in the eggs were lower to their hatchery results. The TL content found in the same development stage in Lockhart Lake was, however, higher to their field results (see Table 4 for TL contents reported from various studies). The difference in TL content between eggs from wild and hatchery-conditioned oysters could be related to the physiological condition of the adult during conditioning. The adults may have been stressed by the hatchery conditioning (temperature variations) as well as by variations in the algae diet. Both stresses may have an effect on gametogenesis. Oysters from the wild may have accumulated more glycogen than those from the hatchery. They might as well have biosynthesized more lipids and consequently supplied more lipid reserves to each egg during vitellogenesis. Frolov & Pankov (1992a) reported that females having gonads with high lipid content produce large larvae with lower ash content. Pretrochophores and veligers collected in the wild were larger than those from the hatchery. However, the size of the trochophores, in relation to their culture environment, was not significantly different and could be explained by a lack of samples. Several factors may affect oocytes development and larval viability during conditioning. Bayne et al. (1978) showed, for instance, that nutritionally stressed mussel females released smaller eggs with less lipid and protein than females in normal conditions. Egg size is strongly correlated to the survival of larvae. This was demonstrated in Mercenaria mercenaria and Argopecten irradians as well (Kraeuter et al. 1982).


Newly-Liberated Larvae

An important decrease in TL content was observed in the hatchery-conditioned oysters when larvae were expelled from the female's paleal cavity. This observation suggests that the lipid content decreased during the veliger stage development. The larvae may have used lipids as energetic reserves to develop complex morphologic structures (e.g., velum, semitransparent shell) whereas they rely on endogenous reserves (Kas'yanov 1984). Helm et al. (1991) and Labarta et al. (1999a) reported similar results at liberation (Table 4). Unfortunately, the TL content for the previous stage (i.e., veliger stage) is, to our knowledge, not available in the literature. Some studies, however, demonstrated that larvae may begin to feed on exogenous sources even before leaving the paleal cavity (Labarta et al. 1999a). The loss of lipids could be related to the fact that newly-liberated larvae remained for a couple of hours without food before they were collected for lipid analysis. They might have been nutritionally stressed and used a part of their lipid reserves. The TAG content decreased during the transition from the veliger stage to a newly-liberated larva. ST and PL contents followed the same trend, but the decrease was less apparent. Newly-liberated larvae might have used sterols and wax esters as a lipid energetic reserve when TAG contents were low. These lipid classes usually decrease once TAG is used for energy (Lee & Hirota 1973, Lee et al. 1974, Frolov & Pankov 1992b). Sasaki (1984) reported, however, that sterols were not catabolized by copepods (Euchaeta japonica) larvae before they were under a 5-6 days starvation.

Our results showed that all lipid contents tend to increase in 8-day old larvae. When larvae are able to feed on exogenous sources, energy in excess is mainly stored as TAG (Gallager et al. 1986). Although there was no significant difference in the lipid content of newly-liberated larvae and 8 day-old larvae, TAG showed the most important increase.

Larvae, which grew in the rearing tank, did not survive more than 10 days. Though the actual causes for mortality were not determined, mass mortality is often related to diseases, variations in the culture environment (Elston et al. 1981), or nutritional deficiencies (Helm et al. 1991). Larval performance is significantly correlated with the lipid content at liberation (Helm et al. 1973). The low survival rate observed in larvae from the hatchery-conditioned oysters may be related to low lipid accumulation.

Larval Quality

The TAG/ST ratio of hatchery-conditioned oysters decreased constantly during the early larval development until larvae were released in the water column by the female. The TAG/ST ratio tends to increase when larvae are able to feed on exogenous sources. In this study, TAG/ST ratios of wild and hatchery early nonfeeding stages were almost similar in spite of their different lipid content. The life-cycle completion according to the culture environment was, however, different. In hatchery-conditioned oysters, larvae did not survive more than 10 days. Larval settlement in the field was, however, abundant (personal observation). This meant that TAG/ST ratios were high enough to support a good physiological quality, because the spat were abundant on the collectors (Burke et al. 2008). Our results, however, suggests that larval quality cannot be only estimated by the TAG/ST ratio. TL contents should also be taken into account.


Our results underline the influence of the conditioning method on the storage of energy and subsequent gametogenesis and spawning events. The genitors will first provide energetic reserves to the eggs, which in turn, will allow larvae to meet their energetic requirements during their development until exogenous food sources are available. Lipid reserves seem to be used in a great amount during the veliger stage development. This observation could not be confirmed by field data. The high larval settlement observed in the field in 2005 (Burke et al. 2008), however, suggests that the lipids supplied to the larvae by the genitors were important enough to sustain the larvae during their development until metamorphosis and settlement.

The variability observed in the lipid content between wild and hatchery-cultured larvae was high for each given stage. A high variability in the lipid content was also observed for a given individual during lipid analysis. The rigor of the analysis could be improved by using a greater number of samples. Still, our study provides an original data set for the study of the physiological quality of early nonfeeding stages, particularly for a species that uses paleal cavity as a brooding chamber. Results from this study showed the limitation of using TAG/ST ratio alone to assess larval quality. The TL content should be taken into account to better estimate the physiological quality of larvae.

The physiological and ecological needs of the European oyster are poorly known. This is particularly true in the context of the colonization of a new habitat. Our field observations showed that larvae may survive off the eastern Canadian coast. However, we do not know whether settlement is constant through the years. It is necessary to monitor this species within a longer time frame. We know that spat may settle on collectors, but we do not know if survival rate is important enough to support the population. Water temperature decreases rapidly between the end of the summer and early fall. Oysters must reach 20 mm to survive their first winter (Newkirk et al. 1995). The survival rate of juveniles after their first winter has never been studied off the eastern Canadian coast. This type of information is necessary to better estimate the recruitment of the European oyster and its aquaculture potential in New Brunswick.


The authors thank D. Wilbur and employees of Hummerfish Aquaculture for their technical support; B. Vercaemer, A. Thompson, S. Roach and B. MacDonald for their assistance in broodstock conditioning and microalgal cultures (Bedford Institute of Oceanography, Dartmouth); M. Robinson and L. Smith for their help on the field and laboratory analysis; and Dr. G. Moreau (Departement de biologic, Universite de Moncton) for his statistical input. Funding for this research was provided by the Department of Fisheries & Oceans and Universite de Moncton.


Abbe, G. R. 1986. A review of some factors that limit oyster recruitment in Chesapeake Bay. Am. Malacol. Bull. 3:59 70.

Akberali, H. B. & E. R. Trueman. 1985. Effects of environmental stress on marine bivalve molluscs. Adv. Mar. Biol. 22:101 198.

Andre, C., P. R. Jonsson & M. Lindegarth. 1993. Predation on settling bivalve larvae by benthic suspension-feeders: the role of hydrodynamics and larval behaviour. Mar. Ecol. Prog. Ser. 97:183-192.

Andrews, J. 1980. A review of introductions of exotic oysters and biological planning for new importations. Mar. Fish. Rev. 42:1-11.

Arellano-Martinez, M., I. S. Racotta, B. P. Ceballs-Vazquez & J. F. Elorduy-Garay. 2004. Biochemical composition, reproductive activity and food availability of the lions paw scallop Nodipecten subnodosus in the Laguna ojo de liebre baja California sur, Mexico. J. Shellfish Res. 23:15-23.

Bataller, E., K. Burke, M. Ouellette & M.-J. Maillet. 2006. Evaluation of spawning period and spat collection of the northernmost population of European oysters (Ostrea edulis L.) on the Canadian Atlantic coast. Can. Tech. Rep. Fish. Aquat. Sci. 2630.

Bartlett, B. R. 1979. Biochemical changes in the Pacific oyster, Crassostrea gigas (Thunberg) during larval development and metamorphosis. Proc. Natl. Shellfish Assoc. 69:202.

Bayne, B. L. 1973. Aspects of the metabolism of Mytilus edulis during starvation. Neth. J. Sea Res. 7:399-441.

Bayne, B. L. 1976. Aspects of reproduction in bivalve molluscs. In: M. Wiley, editor, Estuarine processes. New York: Academic Press. pp. 432-448.

Bayne, B. L., P. A. Gabbott & J. Widdows. 1975. Some effects of stress in the adult on the eggs and larvae of Mytilus edulis L. J. Mar. Biol. Ass. UK 55:675-689.

Bayne, B. L., D. L. Holland, M. N. Moore, D. M. Lowe & J. Widdows. 1978. Further studies on the effects of stress in the adult on the eggs of Mytilus edulis. J. Mar. Biol. Ass. UK58:825-841.

Bayne, B. L. & R. C. Newell. 1983. Physiological energetics of marine molluscs. In: K. M. Wilbur, editor. The mollusca. New York: Academic Press. pp. 407-515.

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 and Reeve). J. Exp. Mar. Biol. Ecol. 79:19-37.

Beukema, J. J., J. Drent & P. J. C. Honkoop. 2001. Maximizing lifetime egg production in a Wadden Sea population of the tellinid bivalve Macoma balthica: a trade-off between immediate and future reproductive outputs. Mar. Ecol. Prog. Ser. 209:119-129.

Brand, A. R., J. D. Paul & J. N. Hoogesteger. 1980. Spat settlement of the scallops Chlamys opercularis (L.) and Pecten maximus (L.) on artificial collectors. J. Mar. Biol. Ass. UK 60:379-390.

Burke, K., E. Bataller, G. Miron, M. Ouellette & R. Tremblay. 2008. Spat collection of a non-native bivalve species (European oyster, Ostrea edulis) off the east Canadian coast. J. Shellfish Res. 27:345-353.

Cook, R. D. 1977. Detection of influential observations in linear regression. Technometrics 19:15-18.

Dame, R. F. 1996. Ecology of marine bivalves: an ecosystem approach. Boca Raton: CRC Press.

Delauney, F., Y. Marty, J. Moal & J.-F. Samain. 1992. Growth and lipid class composition of Pecten maximus (L.) larvae grown under hatchery conditions. J. Exp. Mar. Biol. Ecol. 163:209-219.

Eldredge, L. G. 1994. Introductions of commercially significant aquatic organisms to the Pacific Islands. Noumea, New Caledonia: South Pac. Comm.

Elston, R., L. Leibovitz, D. Relyea & J. Zatial. 1981. Diagnosis of vibriosis in a commercial oyster hatchery epizootic: diagnostic tools and management features. Aquaculture 24:53-62.

Folch, J., M. Lees & G. H. Sloane-Stanlez. 1957. A simple method for the isolation and purification of total lipids from animal tissues. J. Biol. Chem. 226:497-509.

Fraser, A. J., J, C. Gamble & J. R. Sargent. 1988. Changes in lipid content, lipid class composition and fatty acid composition of developing eggs and unfed larvae of cod (Gadus morhua). Mar. Biol. 99:307-314.

Fraser, A. J. 1989. Triacylglycerol content as a condition index for fish, bivalve, and crustacean larvae. Can. J. Fish. Aquat. Sci. 46:1868-1873. Frolov, A. V. & S. L. Pankov. 1992a. The reproduction strategy of oyster Ostrea edulis L. from the biochemical point of view. Comp. Biochem. Physiol. 103b:161-182.

Frolov, A. V. & S. L. Pankov. 1992b. The effect of starvation on the biochemical composition of the rotifer Brachionusplicatilis (Muller). J. Mar. Biol. Ass. UK 72:343-356.

Gabbott, P. A. 1983. Developmental and seasonal metabolic activities in marine molluscs. In: P. W. Hochachka, editor. The mollusca. New York: Academic Press. pp. 165-217.

Gade, G. 1983. Energy metabolism of arthropods and molluscs during environmental and functional anaerobiosis. J. Exp. Zool. 228:415-429.

Gallager, S. M. & R. Mann. 1986. Growth and survival of larvae of Mercenaria mercenaria (L.) and Crassostrea virginica (Gmelin) relative to broodstock conditioning and lipid content of eggs. Aquaculture 56:105-121.

Gallager, S. M., R. Mann & G. C. Sasaki. 1986. Lipid as an index of growth and viability in three species of bivalve larvae. Aquaculture 56:81-103.

Griffond, B., P. Gomot & L. Gomot. 1992. Influence de la temperature sur le deroulement de l'ovogenese chez l'escargot Helix aspersa. J. Therm. Biol. 17:185-190.

Grizel, H. & M. Heral. 1991. Introduction into France of the Japanese oyster (Crassostrea gigas). J. Cons. Int. Explor. Mer. 47:399-403.

Harding, G. C. & A. J. Fraser. 1999. Application of the triacylglycerol/ sterol condition index to the interpretation of larval lobster Homarus americanus distribution in close proximity to Georges Bank, Gulf of Maine. Mar. Ecol. Prog. Ser. 186:239-254.

Haws, M. C., L. Dimichele & S. C. Hand. 1993. Biochemical changes and mortality during metamorphosis of the Eastern Oyster, Crassostrea virginica, and the Pacific oyster, Crassostrea gigas. Mol. Mar. Biol. Biotechnol. 2:202-217.

Helm, M. M., D. L. Holland & R. R. Stephenson. 1973. The effect of supplementary algal feeding of a hatchery breeding stock of Ostrea edulis L. on larval vigour. J. Mar. Biol. Ass. UK 53:673-684.

Helm, M. M., D. L. Holland, S. D. Utting & J. East. 1991. Fatty acid composition of early non-feeding larvae of the flat oyster, Ostrea edulis. J. Mar. Biol. Ass. UK. 691-705.

Hendriks, I. E., L. A. van Duren & P. M. J. Herman. 2003. Effect of dietary polyunsaturated fatty acids on reproductive output and larval growth of bivalves. J. Exp. Mar. Biol. Ecol. 296:199-213.

Holland, D. L. 1978. Lipid reserves and energy metabolism in the larvae of benthic marine invertebrates. In: D. C. Malins & J. R. Sargent, editors. Biochemical and biophysical perspectives in marine biology. New York: Academic Press. pp. 85-123.

Holland, D. L. & P. J. Hannant. 1976. The glycogen content in winter and summer of oysters, Ostrea edulis L. of different ages. J. Cons. Int. Explor. Mer. 36:2402-242.

Holland, D. L. & B. E. Spencer. 1973. Biochemical changes in fed and starved oysters, Ostrea edulis L. during larval development, metamorphosis and early spat growth. J. Mar. Biol. Ass. UK 53:28-298.

Kas'yanov, V. L. 1984. Planktotrophic larvae of bivalve mollusks: morphology, physiology and behaviour. Biol. Morya. 10:17-128.

Korringa, P. 1947. Relations between the moon and periodicity in the breeding of marine animals. EcoL Mon ogr. 17:347-381.

Kraeuter, J. N., M. Castagna & R. Van Dessel. 1982. Egg size and larval survival of Mercenaria mercenaria (L.) and Argopecten irradians (Lamarck). J. Exp. Mar. Biol. Ecol. 56:3-8.

Labarta, U., M. J. Fernhndez-Reiriz & A. Perez-Camacho. 1999a. Larvae of Ostrea edulis L. during starvation: growth, energy and biochemical substrates. Hydrobiologia 405:125-131.

Lannan, J. E. 1980. Broodstock management of Crassostrea gigas. I. Genetic and environmental variation in survival in the larval rearing system. Aquaculture 21:323-336.

Lannan, J. E., A. Robinson & W. P. Breese. 1980. Broodstock management of Crassostrea gigas II. Broodstock conditioning to maximize larval survival. Aquaculture 21:337-345.

Le Dantec, J. 1968. Ecologie et reproduction de l'huitre portugaise Crassostrea angulata Lmk dans le Bassin d'Arcachon et sur la rive gauche de la Gironde. Rev. Tray. Inst. Peches Marit. Paris. 32:1-126.

Lee, R. F. & J. Hirota. 1973. Was ester in tropical zooplankton and nekton and the geographical distribution of wax esters in Omarine copepods. Limnol. Oceanogr. 18:227-239.

Lee, R. F., J. C. Nevenzel & A. G. Lewis. 1974. Lipid changes during life cycle of marine copepod, Euchaeta japonica Marukawa. Lipids 9:891-898.

Lucas, A., L. Chebab-Chalabi & D. Aldana Aranda. 1986. Passage de l'endotrophie h l'exotrophie chez les larves de Mytilus edulis. Oceanol. Acta 9:97-103.

Mann, R. 1979. Some biochemical and physiological aspects of growth and gametogenesis in Crassostrea gigas and Ostrea edulis grown at sustained elevated temperatures. J. Mar. Biol. Ass. UK 59:95-110.

Marteil, L. 1960. Ecologie des huitres du Morbihan Ostrea edulis Linne et Gryphaea angulata Lamarck. Rev. Trav. Inst. Peches Marit. Paris 24:329-446.

Martinez, G. & H. Perez. 2003. Effect of different temperature regimes on reproductive conditioning in the scallop Argopecten purpuratus. Aquaculture 228:153-167.

Mathieu, M. & P. Lubet. 1993. Storage tissue metabolism and reproduction in marine bivalves a brief review. Inv. Reprod. Dev. 23:123-129.

Medcof, J. C. 1961. Trial introduction of European oysters (Ostrea edulis L.) to Canadian east coast. Proc. Nat. Shellfish. Assoc. 50:113-124.

Millar, R. H. & J. M. Scott. 1967. The larva of the oyster Ostrea edulis during starvation. J. Mar. Biol. Ass. UK 47:475-484.

Millican, P. F. & M. M. Helm. 1994. Effects of nutrition on larvae production in the European flat oyster, Ostrea edulis. Aquaculture 123:83-94.

Miron, G., L. J. Walters, R. Tremblay & E. Bourget. 2000. Physiological condition and barnacle larval behaviour: a preliminary look at the relationship between TAG/DNA ratio and larval substratum exploration in Balanus amphitrite. Mar. Ecol. Prog. Ser. 198:303-310.

Navarrete, S. A. & J. C. Castilla. 1990. Barnacle walls as mediators of intertidal mussel recruitment: effects of patch size on the utilization of space. Mar. Ecol. Prog. Ser. 68:113-119.

Newkirk, G. F., B. C. Muise & C. E. Enright. 1995. Culture of the Belon oyster, Ostrea edulis, in Nova Scotia. In: A. Boghen, editor. Coldwater aquaculture in Atlantic Canada. The Canadian Institute for Research on Regional Development, Moncton, pp. 225-253.

Orton, J. H. 1936. Observations and experiments on sex-change in the European oyster (Ostrea edulis). Part V. A simultaneous study of spawning in 1927 in two distinct geographical localities. Mem. Mus. Roy. Hist. Nat. Belg. Ser. 2, Fasc. 3. pp. 997-1056.

Parrish, C. C. 1987. Separation of aquatic lipid classes by Chromarod thin-layer chromatography with measurement by Iatroscan flame ionization detection. J. Fish. Res. Bd. Can. 44:722-731.

Pernet, F., R. Tremblay & E. Bourget. 2003. Biochemical indicator of sea scallop (Plaeopecten magellanieus) quality based on lipid class composition. Part II: Larval growth, competency and settlement. J. Shellfish Res. 22:377-388.

Pernet, F., V. M. Bricelj & C. C. Parrish. 2005. Effect of varying dietary levels of w6 polyunsaturated fatty acids during the early ontogeny of the sea scallop, Placopecten magellanicus. J. Exp. Mar. Biol. Ecol. 327:115-133.

Racotta, I. S., E. Palacios & A. M. Ibarra. 2003. Shrimp larval quality in relation to broodstock condition. Aquaculture 227:107-130.

Rodriguez, J. L., F. J. Sedano, L. O. Garcia-Martin, A. Perez-Camacho & J. L. Sanchez Lopez. 1990. Energy metabolism of newly settled Ostrea edulis spat during metamorphosis. Mar. Biol. 106:109-111.

SAS Institute. 1982. SAS User's Guide: Statistics. Statistical Analysis System Institute, Cary, North Carolina.

Sasaki, G. C. 1984. Biochemical changes associated with embryonic and larval development in the American lobster Homarus americanus Milne Edwards. Ph.D. thesis, Massachusetts Institute of Technology, Cambridge, and Woods Hole Oceanographic Institution, Woods Hole, MA, WHOI-84-8.

Sato, S. 1951. Relationship between the changes in the spawning period and water temperature at Matsushima Bay. Bull. Jpn. Son. Sci. Fish. 7:16-25.

Taylor, A. C. & T. J. Venn. 1979. Seasonal variation in weight and biochemical composition of the tissues of the queen scallop Chlamys opereutaris from the Clyde Sea area. J. Mar. Biol. Ass. UK 59:605-621.

Tocher, D. R., A. J. Fraser, J. R. Sargent & J. C. Gamble. 1985. Lipid class composition during embryonic and early larval development in Atlantic herring (Clupea harengus). Lipids 20:84-89.

Utting, S. D. & B. E. Spencer. 1991. The hatchery culture of bivalve mollusc larvae and juveniles. Ministry of Agriculture, Fisheries and Food Directorate of Fisheries Research.

Waldock, M. J. 1979. The fatty acid composition of the triacylglycerols of oysters, mussels and barnacles. Ph.D. thesis, University College of North Wales.

Walne, P. R. 1974. Observations on the larvae of Ostrea edulis. In: P. R. Walne, editor. Culture of bivalve molluscs: 50 years experience at Conwy. London, England: White Friars Press. pp. 75-111.

Wehrtmann, I. S. 1990. How important are starvation periods in early larval development for survival of Crangon septemspinosa larvae? Mar. Ecol. Prog. Ser. 73:183-190.

Welch, W. R. 1964. The European oyster, Ostrea edulis L., in Maine. Proc. Nat. Shellfish. Assoc. 54:7-23.

Whyte, J. N. C., N. Bourne & C. A. Hodgson. 1989. Influence of algal diets on biochemical composition and energy reserves in Patinopecten yessoensis (Jay) larvae. Aquaculture 78:333-347.

Whyte, J. N. C., N. Bourne & N. G. Ginther. 1990. Biochemical changes during embryogenesis in the rock scallop Crassadoma gigantea. Mar. Biol. 106:239-244.

Zandee, D. I., J. H. Kluytmans, W. Zurburg & H. Pieters. 1980. Seasonal variations in biochemical composition of Mytilus edulis with reference to energy metabolism and gametogenesis. Neth. J. Sea Res. 14:1-29.


(1) Departement de biologie, Universite de Moncton, Moncton, Nouveau-Brunswick, Canada, E1A 3E9; (2) Centre des peches du Golfe, Peches et Oceans Canada, CP 5030, Moncton, Nouveau-Brunswick, Canada, E1C 9B6; (3) Chaire de recherche du Canada en larviculture et production de juveniles en aquaculture, Institut des sciences de lamer, Universite du Quebec a Rimouski, 310 allee des Ursulines, Rimouski, Quebec, Canada, G5L 3A1.

* Corresponding author. E-mail:

Summary of a one-way MANOVA carried out on the mean
lipid content of eggs and larvae from wild European
oysters (Ostrea edulis). Multiple comparisons were
carried out with Tukey-Kramer test. Variable (egg or
larval stage) are presented by increasing order of
lipid content (ng). Nonsignificant differences among
variables are underlined.

Lipid Source of
Classes Variation SS df MS F P

TAG Stage 0.77 2 0.38 1.55 0.236
 (larvae stage) 5.21 21 0.24 3.31 -0.001
 Error 3.52 47 0.07
 Total 9.48 70
ST Stage 0.30 2 0.15 0.32 0.726
 (larvae stage) 9.72 21 0.46 3.28 -0.001
 Error 6.63 47 0.14
 Total 16.65 70
PL Stage 12.22 2 6.11 6.01 0.009
 (larvae stage) 21.35 21 1.01 1.11 0.367
 Error 42.87 47 0.91
 Total 76.40 70
TL Stage 20.02 2 10.01 2.90 0.077
 (larvae stage) 72.37 21 3.44 2.14 0.015
 Error 75.60 47 1.60
 Total 167.78 70

Tukey-Kramer test:

PL content C B#A#

PL content: A#: pretrochhophore; B: trochophore; C: veliger

Note: Nonsignificant differences among
variables are underlined indicated with #.

Summary of a one-way MANOVA carried out on the mean lipid
content of eggs and larvae from hatchery-conditioned European
oysters (Ostrea edulis). Multiple comparisons were carried out
with Tukey-Kramer test. Variable (egg or larval stage) are
presented by increasing order of lipid content (ng). Nonsignificant
differences among variables are underlined.

 Lipid Source of
Classes Variation SS df MS F P

TAG Stage 44.49 4 11.12 13.02 <0.001
 (larvae stage) 18.80 22 3.45 3.45 <0.001
 Error 12.37 50 0.25
 Total 76.38 76
ST Stage 4.75 4 1.18 1.64 0.199
 (larvae stage) 15.92 22 0.72 4.42 <0.001
 Error 8.18 50 0.16
 Total 28.91 76
PL Stage 19.39 4 4.84 2.29 0.092
 (larvae stage) 46.55 22 2.11 2.90 <0.001
 Error 36.52 50 0.73
 Total 102.33 76
TL Stage 68.89 4 17.22 3.59 0.021
 (larvae stage) 105.43 22 4.79 3.49 <0.001
 Error 68.71 50 1.37
 Total 244.02 76

Tukey-Kramer test:

TAG content A# B# C# E# D#
TL content C# A# B# E# D#
PL content: A: pretrochophore D: newly-liberated larvae
B: trochophore E: 8 day-old larvae
C: veliger

Note: Nonsignificant differences among variables are underlined
indicated with #.


Area (mean [+ or -] SD) of pretrochophore, trochophore and veliger
stages sampled from wild and hatchery-conditioned European
oysters (Ostrea edulis).

 Wild ([micro] Hatchery ([micro]
 [m.sup.2]) [m.sup.2])

"White sick" 10,959 [+ or -] 1,286 9,110 [+ or -] 861
n 500 300
"Grey sick" 15,839 [+ or -] 1,583 15,416 f 2,971
n 300 300
"Black sick" 23,830 [+ or -] 3,020 23,108 [+ or -] 1,906
n 500 200

TABLE 4. Total lipid (TL) content comparison for various
developmental stages of European oysters (Ostrea edulis).

 Wild Hatchery

Pre-trochophore 94.61 [+ or -] 23.87 44.04 [+ or -] 21.16
Trochophore 88.53 [+ or -] 9.41 47.72 [+ or -] 14.64
Veliger 72.75 [+ or -] 16.14 51.81 [+ or -] 9.78
 larvae n/a 22.30 [+ or -] 9.23

 Wild (Helm et Hatchery (Helm
 al. 1991) et al. 1991)

Pre-trochophore 71 [+ or -] 7 79 (non published data)
Trochophore 69 [+ or -] 11 n/a
Veliger 64 [+ or -] 23 n/a
Newly-liberated 29-33 [+ or -] 3
 larvae n/a 27 (Labarta et al. 1999a)
COPYRIGHT 2008 National Shellfisheries Association, Inc.
No portion of this article can be reproduced without the express written permission from the copyright holder.
Copyright 2008 Gale, Cengage Learning. All rights reserved.

 Reader Opinion




Article Details
Printer friendly Cite/link Email Feedback
Author:Burke, Kevin; Bataller, Erick; Miron, Gilles; Ouellette, Marc; Tremblay, Rejean
Publication:Journal of Shellfish Research
Article Type:Report
Geographic Code:1CANA
Date:Aug 1, 2008
Previous Article:Harvest history and current densities of the pearl oyster Pinctada mazatlanica (Bivalvia: pteriidae) in Las Perlas and Coiba Archipelagos, Panama.
Next Article:Prospective culture of the Cortez oyster Crassostrea corteziensis from northwestern Mexico: growth, gametogenic activity, and condition index.

Related Articles
Relationship of amebocytes and terrestrial elements to adult shell deposition in eastern oysters.
Effects of salinity on growth and survival of silver-lip pearl oyster, Pinctada maxima, spat.
Microscopic observations of larval Ostrea circumpicta (bivalve: ostreidae) in brood chambers.
Age and growth of wild suminoe (Crassostrea ariakensis, Fugita 1913) and Pacific (C. gigas, Thunberg 1793) oysters from Laizhou Bay, China.
The limits of morphometric features for the identification of black-lip pearl oyster (Pinctada margaritifera) larvae.
Crassostrea ariakensis in Chesapeake Bay: growth, disease and mortality in shallow subtidal environments.
Responses of Crassostrea virginica (Gmelin) and C. ariakensis (Fujita) to bloom-forming phytoplankton including ichthyotoxic Karlodinium veneficum...
Prospective culture of the Cortez oyster Crassostrea corteziensis from northwestern Mexico: growth, gametogenic activity, and condition index.
A comparison of the macrofaunal communities inhabiting a Crassostrea virginica oyster reef and oyster aquaculture gear in Indian River Bay, Delaware.
Prospect of oyster culture in Pakistan: pathology assessment of two commercially important oyster species.

Terms of use | Copyright © 2014 Farlex, Inc. | Feedback | For webmasters