Aquacult Int (2013) 21:1065–1076 DOI 10.1007/s10499-012-9612-7
Embryonic development and influence of egg density on early veliger larvae and effects of dietary microalgae on growth of brown mussel Perna perna (L. 1758) larvae under laboratory conditions Lahoussine Aarab • Alejandro Pe´rez-Camacho • Marı´a del Pino Viera-Toledo • Gercende Courtois de Vic¸ose Hipo´lito Ferna´ndez-Palacios • Lucia Molina
•
Received: 2 July 2012 / Accepted: 24 October 2012 / Published online: 6 November 2012 Ó Springer Science+Business Media Dordrecht 2012
Abstract The effect of egg density on embryonic development and larval quality as well as the lipid and fatty acid contents (eicosapentaenoic acid, EPA; docosahexaenoic acid, DHA) of cultured microalgae fed to Perna perna larvae was studied under controlled conditions to provide information needed for development of an experimental hatchery. Embryonic development followed the common sequence exhibited by other bivalves. D-larva stage was attained 40–44 h post-fertilisation at 21 ± 1 °C. The umbo-stage was reached in 11 days, and pediveliger larvae were observed 26 days post-fertilisation. Low egg density (range 20–100 eggs cm-2) produced high proportions of normal D-larvae. Larval development showed two growth phases: 1st—the mixotrophic stage and 2nd—the exotrophic stage where the composition of diets had significant effects on larval growth with higher rates in larvae fed with the mixed microalgae (Isochrysis galbana ? Chaetoceros calcitrans, I. galbana ? Phaeodactylum tricornutum and I. galbana ? Skeletonema costatum) in comparison with the monospecific diet (I. galbana). Fatty acid analysis showed that larval growth and survival were strongly influenced by proportions of dietary DHA and EPA. These results indicate that DHA and EPA are the key factors in determining larval performance, considerably more than the total amount of other fatty acids. Keywords Brown mussel Egg densities Growth Hatchery Larvae Microalgae Perna perna Survival Abbreviations EPA Eicosapentaenoic acid DHA Docosahexaenoic acid ARA Arachidonic acid ANOVA Analysis of variance L. Aarab (&) M. P. Viera-Toledo G. C. de Vic¸ose H. Ferna´ndez-Palacios L. Molina Grupo de Investigacio´n en Acuicultura, ULPGC, ICCM, P. O. Box 56, 35200 Telde, Canary Islands, Spain e-mail:
[email protected] A. Pe´rez-Camacho Centro Oceanogra´fico de La Corun˜a, IEO, Muelle de Animas, s/n, 18001 La Corun˜a, Spain
123
1066
SFA MUFA PUFA
Aquacult Int (2013) 21:1065–1076
Saturated fatty acids Monounsaturated fatty acids Polyunsaturated fatty acids
Introduction Mussel farming is centuries old and is still actively practiced in many forms around the world with the most common species cultured being of the genus Mytilus and Perna. In 2008, 1.62 million tonnes of mussels were produced worldwide (FAO 2010). In 2007, Spain was the second largest world producer of mussels (250,000 t/year), China being first (450,000 t/year; Figueras 2007). Mussel farming throughout the world, including the leading countries, relies entirely on natural stocks for procurement of seed. The unpredictability and scarcity of wild seed due to various environmental and biological factors (Marshall et al. 2010) and also due to strong restrictions on collection of wild mussel seed enforced by strict environmental and wild life protection laws (Domı´nguez et al. 2010) are becoming a concern for mussel producers. Mussel hatcheries could provide an alternative source of mussel spat to avoid environmental constraints (Spencer 2002). In Spain and the Iberian Peninsula, the species with the highest economic interest is the Mediterranean mussel Mytilus galloprovincialis (Figueras 1989), while the brown mussel Perna perna presents an important conservation interest in the Canary Islands (Carrillo et al. 1992). Overexploitation of P. perna has lead to mussel collection banning by the Canary Islands Government (Decree 134/1986, September 12), based on the fact that unreliable natural recruitment rates do not allow development of controlled and sustainable fisheries of this resource. Controlled hatchery production of juvenile mussels would allow a faster recovery of wild populations and promote development of commercial culture of this species (Pere´z-Camacho et al. 1995; Beaumont and Hoare 2003). However, significant production problems of juvenile still remain; the most stringent limitations include the following: rearing bivalves to post-larval stages in order to complete the life cycle and the mass-production of microalgae for feeding and optimising environmental parameters during larval stages (Robert and Gerard 1999). The major objective in hatchery production is to improve larval survival by maximising larval growth and success at metamorphosis of the newly settled seed. Despite many studies on mussel larval development carried out at the laboratory scale (e.g. Bayne 1965; Ve´lez and Martı´nez 1967; Beaumont and Budd 1982; Sprung and Bayne 1984; Leonardos and Lucas 2000a; Laxmilatha et al. 2011), optimisation of larvae nutrition and culture is required to ensure fast growth rates that would be needed for mass seed production (Galley et al. 2009; Ragg et al. 2010). The aim of this study was to investigate embryological development and the effects of egg density on early veliger larvae, and the effect of different diets on survival and growth of the brown mussel P. perna larvae reared under experimental conditions.
Materials and methods Broodstock maintenance and spawning of Perna perna Fifty adult P. perna, with shell lengths of 55.13 ± 18.03 mm, were collected in April 2010, from natural populations along ‘‘Rincon de Los Morteros’’, in Fuerteventura Island.
123
Aquacult Int (2013) 21:1065–1076
1067
These were transported to the mussel hatchery of the Instituto Canario de Ciencias Marinas (Canary Islands, Spain). Prior to the experiment, broodstock was kept for 1 week in aerated seawater in 50-l tanks and fed daily with 3 9 105 cells ml-1 of Isochrysis galbana and Chetoceros calcitrans (1:1 based on cell counts). Water temperature was kept at 21 ± 1 °C during the experiment. The shells were carefully cleaned by scrubbing and rinsing them in filtered seawater. Spawning was induced by thermal shock, attained by transferring the mussels alternatively between trays with water temperatures at 16 or 25 °C at hourly intervals until spawning (about 3 h). Specimens responding to the thermal stimulation were individually placed in a 100-ml beaker for release of sperm or eggs. Males and females were identified at the time of gamete release. Oocytes and spermatozoids were mixed in a 10-l cylinder at a ratio of 5–10 spermatozoids/egg (Beaumont et al. 2004), and the resulting fertilised eggs remained untouched in the 10-l cylinders until the appearance of the first polar body. Seawater (21 °C) was filtered through a 1-lm Millipore cartridge filter and UV treated. Fertilised eggs were then sieved on a nylon screen (25-lm mesh) to eliminate excess sperm and placed into flat-bottomed glass dishes (Experiment 1: to determine optimum egg density) or in 50-l tanks at a density adjusted to 50 eggs cm-2 (Experiment 2: to study embryo development). Cultures were maintained at 21 ± 1 °C without aeration. All experiments were conducted with three replicates per treatment. Embryonic development and egg density effect on the quality of early veliger Experiment 1 To study the effect of egg density, fertilised eggs were distributed into glass dishes containing 50 ml of fresh 1-lm filtered and UV sterilised seawater at densities of 20, 50, 100, 200 or 400 eggs cm-2. Larvae were inspected after approximately 48 h and assessed, using an optical microscope, for the percentage that showed normal morphology (normality) as opposed to abnormal ones from random samples of 100 larvae. Experiment 2 To study embryonic development of P. Perna, 100 ml of seawater containing embryos was sampled on a 40-lm-mesh nylon screen every hour, and 100 embryos were observed under optical microscope. The morphological characteristics of each embryonic stage observed were compared to those of Perna viridis described by Sreenivasan et al. (1988) due to the limited number of descriptive studies on P. perna embryonic development. Differentiation between two stages was established when more than 50 % of the embryos from the sample reached a following stage. Effect of different diets on growth and survival of larvae Experiment 3 44 h after hatching early veliger larvae were transferred to 12 10-l tanks at a density of 10 larvae ml-1 according to Galley et al. (2009). Larvae were reared in a closed culture system with UV-treated, 1-lm filtered seawater at 21 ± 1 °C, gently aerated via air diffusers and entirely renewed every 3 days. In the first 11 days, microalgae were fed once a day at a concentration of 4 9 104 cells ml-1 and increased gradually up to 12 9 104 cells ml-1 twice a day (Beduschi et al. 2009). The experiment was completed when larval settlement and metamorphosis were observed. For each feeding experiment, four microalgal diets, further described, were compared, with three replicates per treatment.
123
1068
Aquacult Int (2013) 21:1065–1076
Microalgal culture and lipid content determination Microalgae used in the feeding experiments were cultured onsite from initial cultures purchased from different collections. Four species were cultured: Isochrysis galbana (BNA-40-002) and Phaeodactylum tricornutum from Instituto Tecnolo´gico de Canarias ITC (Spain), Chaetoceros calcitrans (provided by the Instituto Galego de Formacio´n en Acuicultura IGAFA (Spain) and Skeletonema costatum (CCAP 1077/1B) from Dunstaffnage Marine Laboratory (UK). All species were grown in a series of 40-l bags with 1-lm filtered and UV-treated seawater at a salinity of 36 % enriched with F2 medium for I. galbana and (F2 ? Silicate) medium for the diatoms C. calcitrans, S. costatum and P. tricornutum. All cultures received continuous aeration and were grown under a 24 h light regime (5,000 ± 850 lux) at 19 ± 1 °C. Larvae were fed a controlled diet of I. galbana and a combination of the flagellate I. galbana and the diatoms C. calcitrans, S. costatum and P. tricornutum. The diets used were the following: 1. 2. 3. 4.
(Is): Isochrysis galbana, (Is ? Ch): a 1:1 (based on cell counts) mixture of I. galbana and C. calcitrans, (Is ? Sk): a 1:1 (based on cell counts) mixture of I. galbana and S. costatum, (Is ? Ph): a 1:1 (based on cell counts) mixture of I. galbana and P. tricornutum.
Microalgae used in this study exhibited a similar size in their exponential phase of growth (40–45 lm3 equivalent to 4.2–4.4 lm diameter) as well as similar dry individual weight (18–20 pg: Robert et al. 2004). Samples of algae were harvested in the early stationary phase, concentrated by centrifugation and stored at -80 °C, and lipid was extracted as described by Folch et al. (1957). Fatty acid methyl esters were obtained by transmethylation of crude lipids as described by Christie (1982) purified on NH2 silica (Sep-pak; Waters) and separated and quantified in a Shimadzu gas chromatograph (GC-14A; Shimadzu, Kyoto, Japan) under the conditions described in Izquierdo et al. (1990) and identified by comparison with previously characterised standards and GLC–MS (Gas–Liquid Chromatographic-Mass Spectrometric). Sampling and husbandry of larvae At 3-day intervals during the culture period, larvae were sieved on a 60-lm up to 150-lm-mesh screen and inspected, the tanks were cleaned and the seawater replaced with no antibiotic addition. Shell length was measured as the greatest dimension in a line parallel to the hinge (anterior–posterior axis), and shell height was measured as the perpendicular dorso–ventral axis, according to Bayne (1976). Since the growth axis of the mussel is along the dorso-ventral, the growth of the larvae was deduced from the dorsoventral measurements. Mean values of larval growth were acquired from 30 measurements per tank using image analysis technique (Image-Pro Plus version 5.0.1 package: Media Cybernetics L.P., Carlsbad, USA). Survival rates during the culture period were measured when larvae reached the umbo-veliger (11D) and pediveliger larval stages and prior to settlement and metamorphosis assays (26D). Data analysis Statistical analysis was facilitated using SPSS 15.0 for Windows (SPSS. Inc., Chicago, IL, USA). Results were checked for normality and homogeneity of variances using the
123
Aquacult Int (2013) 21:1065–1076
1069
Table 1 Percentage of normal P. perna veliger larvae cultured at different egg densities (eggs cm-2), 48 h after fertilisation at 21 ± 1 °C Egg density (eggs cm-2)
20 (n = 100)
50 (n = 100)
100 (n = 100)
200 (n = 100)
400 (n = 100)
Mean (%) (SD)
76.60 (4.85)a
78.73 (4.61)a
66.20 (7.08)a
46.53 (4.07)b
40.53 (6.31)b
Mean values with different superscript letters are significantly different (P \ 0.05). Standard deviation in brackets (SD). n number of samples
Kolmogorov–Smirnov and Levene’s test, respectively, and analysed using a one-way ANOVA, and means were compared by Duncan’s test (P \ 0.05).
Results Egg density effect on the quality of early veliger (Experiment 1) Table 1 shows the percentage of normal morphological development of veliger P. perna, 48 h after fertilisation at different egg densities. The density of P. perna fertilised eggs had a significant effect on development of normal veliger larvae. The percentage of larval normality was not significantly different between densities of 20, 50 and 100 eggs cm-2 (76.60, 78.73 and 66.20 %, respectively). However, concentrations of 200 and 400 eggs cm-2 gave significantly lower rates of normal veliger larvae (46.53 and 40.53 %, respectively). Embryonic development (Experiment 2) Oocytes were 40–60 lm diameter spherical cells with a thin vitteline coat (Fig. 1a). Table 2 shows the different embryonic stages of P. perna and the times at which they occurred at 21 ± 1 °C. Appearance of the first polar body 15 min after oocyte and sperm contact (t0 ? 15 min) is a visual witness of fertilisation. The first segmentation cleavage occurred 40 min after fertilisation (t0 ? 40 min) and results in two blastomeres. The second and third segmentation cleavages occurred between t0 ? 1 h 30 min and t0 ? 2 h. Successive cleavages result in a ciliated morula at (t0 ? 2h 30 min), followed by a ciliated blastula, 2 h later (t0 ? 4h 30 min), and gastrula, 2 h later (t0 ? 6h 30 min). Trochophores were observed actively swimming after 18 h (t0 ? 18 h), and D-larva (Fig. 1b) appeared in the cultures 40–44 h after fertilisation. From t0 ? 40 h, the calcified shell valves enlarge until they enclose the whole soft body. Embryos were observed to rotate continuously at all times. Fatty acid composition of algae and effect of different diets on survival and growth of larvae (Experiment 3) Figure 2 shows the shell height of P. perna larvae fed on different microalgal diets. At 48 h after fertilisation, the D-larvae, with an initial size of 88.22 9 66.54 lm (Fig. 1b), had a developed digestive system, consisting of a mouth, a foregut and a digestive gland. The mantle cavity was evident at this stage of development. Eleven-day-old larvae, which became oval-shaped, were umbonate with dimensions between 118.43 9 96.08 (Is) and 126.93 9 100.82 lm (Is ? Sk) (Fig. 1c). Emergence of the umbonate phase began initially at 8 days and dominated the stage from day 14 onwards
123
1070
Aquacult Int (2013) 21:1065–1076
Fig. 1 Perna perna: larval development: a Fertilised egg; b ‘‘D’’ stage; c Umbo; d Pediveliger Table 2 Timing of the early embryonic development of P. perna reared at 21 ± 1 °C
Differentiation between two stages was established when more than 50 % of the embryos from the sample of 100 embryos reached a following stage
Stage
Time after fertilisation
Egg
t0
1st polar body
t0 ? 15 min
2-cell
t0 ? 40 min
4-cell
t0 ? 1 h 30 min
8-cell
t0 ? 2 h
Morula
t0 ? 2 h 30 min
Blastula
t0 ? 4 h 30 min
Gastrula
t0 ? 6 h 30 min
Trochophore
t0 ? 18 h
D-Larvae
t0 ? 40–t0 ? 44 h
and was followed by eyed larvae characterised by the presence of a black rounded spot below the food mass (20–23-day-old). Twenty-six days after fertilisation, they reached the pediveliger stage, at sizes between 250.14 9 222.45 (Is) and 278.83 9 259.52 lm (Is ? Sk) (Fig. 1d), crawling and swimming for short intervals. With the formation of the foot, disappearance of ciliated velum and appearance of gill filaments, the larvae tended to settle. The relationship between shell length and shell height of the larvae was linear (Fig. 3). The linear relationship is given by the following equation: Y ¼ 0:923X 15:22; R2 ¼ 0:98574;
123
Aquacult Int (2013) 21:1065–1076
1071
Fig. 2 Shell height (lm) of P. perna larvae fed on different microalgal diets during larval development at 21 ± 1 °C. On day 2, the shell heights are equal for all treatments because one sample was measured and then divided over tanks. Mean values with different superscript letters are significantly different (P \ 0.05)
Fig. 3 Relationship between shell length and height of the P. perna larvae
where Y is the shell height (dorso-ventral axis) and X is the shell length (Anterior-posterior axis). Total fatty acids were isolated and identified in the four algal species tested with significant differences in lipid content and fatty acid generally detected between microalgae (Table 3). I. galbana showed significantly higher overall total lipid content and total fatty acid (mg/g of lipid) (58.79 and 200.58 % respectively), while total fatty acid per g of algae dry weight was higher in diatoms P. tricornutum (441 %). However, the different fatty acid classes (SFA, MUFA and PUFA) did not show large variations (24–36, 28–41 and 33–40 %, respectively). However, docosahexaenoic acid (DHA, 22:6n-3) was significantly highest in I. galbana (9.30 %), whereas eicosapentaenoic acid (EPA, 20:5n-3) was higher in the diatoms C. calcitrans (18.65 %), P. tricornutum (23.37 %) and S. costatum (20.60 %) than in I. galbana (0.81 %). Arachidonic acid (ARA, 20:4n-6) was significantly higher in C. calcitrans (4.08 %), but there were no significant differences in total n-3 PUFA among the four algae. On the contrary, total n-6 PUFA levels were significantly higher in diatoms than in I. galbana. The n-6 LC-PUFA was significantly higher in C. calcitrans, whereas I. galbana was significantly higher in n-3 LC-PUFA (10.27 %). Larvae fed on different diets showed similar survival and growth rates during the first 11 days of rearing (Table 4) corresponding to the first growth phase. However, after 14 days of culture, lowest mortality and highest growth rates were observed with significant differences among treatments, corresponding to the second growth phase (Fig. 2). Is ? Sk and Is ? Ph were the best diets leading to high survival (88.54 and 85.42 %, respectively) and average growth of 10.13 and 10.15 lm day-1, respectively, during the second growth phase (Table 4) although in certain days without significant differences with those fed Is ? Ch (Fig. 2). In contrast, larvae fed exclusively I. galbana led to a
123
1072
Aquacult Int (2013) 21:1065–1076
Table 3 Fatty acid profile of cultured microalgae I. galbana (n = 3) 20:4n-6 (ARA) 20:5n-3 (EPA) 22:6n-3 (DHA)
C. calcitrans (n = 3) 4.08 (0.07)a
1.09 (0.08)b
1.05 (0.07)b
d
c
a
20.60 (0.26)b
b
1.90 (0.06)b
c
24.05 (0.17)c
ab
41.11 (0.67)a
a
33.40 (0.52)a
0.81 (0.32)
18.65 (0.88)
a
9.30 (2.77)
1.25 (0.16)
36.18 (2.11) 28.30 (0.97)
a
35.39 (3.09)
b
Total FA (mg/g of lipid) Total FA (mg/g of algae dry weight)
21.46 (1.11)
b
4.22 (0.07)
a
1.30 (0.16)
a
17.69 (2.05) a
200.58 (16.17)
b
341.88 (12.09)
1.39 (0.11)b
b
2.52 (0.08)b
2.81 (0.74)
c
58.79 (6.81)
23.88 (0.11)a
b
1.28 (0.07)
b
10.27 (3.10)
a
27.47 (0.69)
a
1.16 (0.91)
4.73 (0.75)ab
6.06 (0.11)
a
25.11 (2.50)
% Lipid (dry weight)
a
8.10 (2.07)
a
n-3 LC-PUFA
40.06 (0.68)
a
1.58 (0.94)
n-6 LC-PUFA
33.60 (0.23)
a
39.07 (4.06)
n-3 PUFA
25.94 (0.46)
b
28.47 (1.71)
n-6 PUFA
2.30 (0.65)
a
b
PUFA
23.37 (0.11)
b
31.71 (3.35)
MUFA
S. costatum (n = 3)
0.42 (0.03)c
b
SFA
P. tricornutum (n = 3)
bc
36.79 (2.77)b
b
130.97 (17.01)b
26.54 (1.37)
c
46.73 (2.49)
116.94 (0.16) c
265.13 (16.68)
a
441.16 (22.10)
355.26 (19.50)b
Values are percentages of total fatty acid of algae Different letters in same row indicate significant differences (P \ 0.05). Standard deviation in brackets (SD). n number of samples
Table 4 Mean shell height, survival and growth rate (SD) of P. perna larvae fed on different mono- or pluri-specific diets on days 11 and 26 Diet
Height (lm) (D11)
Survival (%) (D11)
Growth (lm day-1) (D2-D11)
Height (lm) (D26)
Survival (%) (D26)
Growth (lm day-1) (D11-D26)
Is
118.66 (9.05)a
98.91 (0.43)a
3.38 (1.25)a
250.14 (25.39)c
72.63 (12.29)d
8.76 (1.63)c
Is ? Ch (n = 90)
124.93 (11.02)a
99.32 (0.17)a
4.08 (1.08)a
266.29 (21.41)b
80.16 (7.38)c
9.42 (1.50)b
Is ? Ph (n = 90)
123.40 (13.87)a
99.13 (0.28)a
3.91 (1.86)a
275.67 (32.60)ab
85.42 (5.91)b
10.15 (1.81)a
Is ? Sk (n = 90)
126.93 (15.75)a
99.82 (0.10)a
4.30 (1.73)a
278.83 (31.15)a
88.54 (7.26)a
10.13 (2.06)a
(n = 90)
Mean values with different superscript letters are significantly different (P \ 0.05). Initial larval shell height on D2 = 88.22 lm (8.94). n number of samples
significant low growth (8.76 lm day-1) and low survival on day 26 (72.63 %) and were characterised by high intertank variability (12.29 %). The same results were also found when growth rates were analysed (Table 4).
Discussion Density of eggs can be measured relative to the volume of a container (eggs ml-1) or relative to the surface area of the bottom of the container (eggs cm-2). Because eggs of P. perna settle quickly to the bottom, surface area is a critical factor in the early stage of
123
Aquacult Int (2013) 21:1065–1076
1073
development while the embryos are non-motile, whereas the volume of the container is a recommended parameter for motile embryos (Galley et al. 2009). Few studies on optimum egg incubation densities for bivalves have used container volume: 30 eggs ml-1 for Tivela mactroides (Silberfeld and Gros 2006), Argopecten nucleus and Nodipecten nodosus (Velasco et al. 2007), and 20 eggs ml-1 for symbiotic calm Codakia orbicularis (Gros et al. 1997) and for basket cockle Clinocardium nuttallii (Liu et al. 2008). Other studies used the bottom surface for the distribution of eggs and showed good results for low densities: Pecten maximus at 7 eggs cm-2, and also, a concentration of 700 eggs cm-2 gave acceptable results for the number of normal larvae (Gruffydd and Beaumont, 1970). Pere´z-Camacho (1977) showed that the percentage of normal larvae is 84–97 % in the case of incubation densities of 1,000–3,000 eggs cm-2 for Venerupis pullastra. For mussels, Sprung and Bayne (1984) advocated low concentrations of embryos (20 eggs cm-2) for Mytilus edulis. However, Galley et al. (2009) produced large proportions of normal larvae ([80 %) using mean densities of 200 M. edulis eggs per cm2. Also, high concentrations of eggs and embryos (1,000 eggs cm-2) are still recommended by the studies of Helm and Bourne (2004). In our study, the low concentrations of P. perna eggs (20, 50 and 100 eggs cm-2) produced high proportions of normal larvae, and these results indicate that P. perna embryos developed best up to normal D-larvae in the egg density range of 20–100 cm-2. At high egg concentrations (200 and 400 eggs cm-2), production of normal larvae was significantly reduced. These results were consistent with the study of Sprung and Bayne (1984), which adopted an initial concentration of 20 eggs cm-2 in M. edulis. The studies cited have shown large variations in the proportions of normal larvae produced by different densities of bivalve eggs. These variations are probably due to physiological conditions of breeding (Loosanoff and Davis 1963), genetic variation (Beaumont and Budd 1983) and (Galley et al. 2009) or water quality (Utting and Helm 1985). Many studies have investigated various aspects of embryonic and larval development in some bivalve species (e.g. Madrones-Ladja 1997; Sreenivasan et al. 1988; Moue¨za et al. 2006; Velasco et al. 2007; Da Costa et al. 2008; Liu et al. 2008; Ruiz et al. 2008). The embryonic development of P. perna has a similar development compared to previously studied bivalves P. viridis (Sreenivasan et al. 1988), Mytilus galloprovincialis (Ruiz et al. 2008) and Ensis arcuatus (Da Costa et al. 2008), although it was slightly faster. Gastrulae were observed 6–7 h after fertilisation, trochophores 11 h later, and fully developed veligers appeared by 40–44 h at 21 °C while it took M. galloprovincialis around 45 h at 17 °C to reach its veliger stage. The nutritional value of algae for most bivalves is determined by essential fatty acids eicosapentaenoic acid (EPA) 20:5(n-3) and docosahexaenoic acid (DHA) 22:6(n-3) (Lane 1989). These polyunsaturated fatty acids (PUFAs) are essential because most bivalve species are unable to produce them from shorter chain precursors (Ackman 1983; Chu and Greaves 1991; Delaunay et al. 1993). Algae differ in their lipid content (Brown et al. 2001; Eglinton et al. 1996), and more specifically, PUFA content (Leonardos and Lucas 2000a; Napolitano et al. 1990). Fatty acid profiles of cultured microalgae, used to feed larvae of P. perna, were comparable to those previously reported (Napolitano et al. 1990; Brown et al. 1997; FernandezReiriz et al. 1998; Leonardos and Lucas 2000b; Pettersen et al. 2010). However, biochemical composition has been shown to vary considerably, depending on culture conditions (Thompson et al. 1996). This study clearly shows that algal composition of the diet can significantly affect performance of hatchery reared brown mussel larvae. Our experiments showed that higher
123
1074
Aquacult Int (2013) 21:1065–1076
growth was observed in larvae fed with the three mixed diets (Is ? Ch, Is ? Ph and Is ? Sk) compared with the monospecific diet (I. galbana), presumably because a mixed algal diet increases the chances of achieving a balanced diet and provides a greater diversity of biochemical constituents to assure most nutritional rations for growth and survival (Widdows 1991; Muller-Feuga et al. 2003; Rico-Villa et al. 2006; Galley et al. 2009). The diatoms C. calcitrans, P. tricornutum and S. costatum had significant effects on increasing growth and survival of larvae. Variation between growth rates and diet was clearly evident when M. galloprovincialis larvae were reared on a diet high in C. calcitrans (Pettersen et al. 2010). Rico-Villa et al. (2006) and Ragg et al. (2010) consider lipid quality rather than quantity to be of prime importance in determining feed quality, particularly the relative amount of the n-3 fatty acids EPA (C20:5n-3) and DHA (C22:6n-3). As well Knauer and Southgate (1999) show that the nutritional value of algae to most bivalve larvae is partially determined by the amounts of the essential fatty acids EPA and DHA they contain. In this study, the high EPA concentrations observed in diatoms C. calcitrans, P. tricornutum and S. costatum (18.65, 23.37 and 20.60 %, respectively) were consistent with the study of Delaunay et al. (1993), Rico-Villa et al. (2006) and Pettersen et al. (2010). I. galbana contained very low level of EPA (0.81 %) while the levels of DHA were lower in diatoms (between 1.25 and 2.30 %) than in the flagellate I. galbana (9.30 %), consistent with levels of DHA reported by Delaunay et al. (1993), Rico-Villa et al. (2006) and Pettersen et al. (2010). DHA is required for development of young bivalves (Enright et al. 1986), but is only present at very low levels in diatoms. However, I. galbana contains relatively high DHA levels 9–10 %, providing an explanation for the complementary benefit of combined diatoms (C. calcitrans, P. tricornutum and S. costatum)/I. galbana diet. This may explain higher growth rates of P. perna larvae fed mixed diets rather than monospecific diet (I. galbana). Robert et al. (2001) showed that the use of monospecific diets in the feeding stage indicate that these are not beneficial for bivalve larvae while mixed diets have a positive effect on growth and mortality compared to diets consisting of only one algal species. Galley et al. (2009) also observed that M. edulis larvae grew faster when fed with a mixed diet of C. calcitrans and I. galbana in comparison with the monospecific diets of these algal species. Acknowledgments This study was funded by JACUMAR (Junta Nacional Asesora de Cultivos Marinos) within the project ‘‘Culture of mussels: the expansion and sustainability’’. We would like to thank Professor M.S. Izquierdo for correcting the manuscript, and Mrs S. Merbah (PhD Student in Houari Boumediene University in Algeria) and Mr F. Otero (PhD Student in Las Palmas de Gran Canaria University in Spain) for their assistance in the establishment of the mussel hatchery. We would also like to thank two anonymous reviewers for their helpful comments to improve the manuscript.
References Ackman RG (1983) Fatty acid metabolism of bivalves. In: Pruder GD, Langdon CJ, Conklin DE (eds) Louisiana State University, Baton Rouge, LA, USA, pp 358–376 Bayne BL (1965) Growth and the delay of metamorphosis of the larvae of Mytilus edulis (L.). Ophelia 2:1–47 Bayne BL (1976) Marine mussels: their ecology and physiology. Cambridge University Press, London Beaumont AR, Budd MD (1982) Delayed growth of mussel (Mytilus edulis) and scalopp (Pecten maximus) veligers at low temperatures. Mar Biol 71:97–100 Beaumont AR, Budd MD (1983) Effects of self-fertilisation and other factors on the early development of the scallop Pecten maximus. Mar Biol 76:285–289 Beaumont AR, Hoare K (2003) Biotechnology and genetics in fisheries and aquaculture. Chapman & Hall, London
123
Aquacult Int (2013) 21:1065–1076
1075
Beaumont A, Turner G, Word A, Skibinski D (2004) Hybridisations between Mytilus edulis and Mytilus galloprovincialis and performance of pure species and hybrid veliger larvae at different temperatures. J Exp Mar Biol Ecol 302:177–188 Beduschi P, Rodriques de Melo CM, Ferreira JF (2009) The influence of techniques of larvae rearing and seed collectors on the survival rate and recovery efficiency of the brown mussel Perna perna (L.) in laboratory. Braz Arch Biol Tech 52(1):145–152 Brown MR, Jeffry SW, Volkman JK, Dunstan GA (1997) Nutritional proprieties of microalgae for mariculture. Aquaculture 151:315–331 Brown MR, Mccausland MA, Kowalski K (2001) The nutritional value of four Australian microalgal strains fed to pacific oyster Crassostrea gigas spat. Aquaculture 165:281–293 Carrillo M, Bacallado JJ, Cruz T (1992) Primeros datos sobre el mejilllo´n Perna Perna (Linnaeus, 1758) en la costa de Fuerteventura (islas Canarias). In: Bacallado JJ, Barquı´n J (eds) Actas del V Simposio Ibe´rico de Estudios del Bentos Marino. Universidad de La Laguna, Museo Insular de Ciencias Naturales and Universidad Polite´cnica de Las Palmas, La Laguna, pp 411–422 Christie WW (1982) Lipid analysis. Pergamon, Oxford Chu FE, Greaves J (1991) Metabolism of palmitic, linoleic, and linolenic acids in adult oysters, Crassostrea virginica. Mar Biol 110:229–236 Da Costa F, Darriba S, Martı´nez-Patin˜o D (2008) Embryonic and larval development of Ensis arcuatus (Jeffereys, 1865) (Bivalvia: Pharidae). J Molluscan Stud 74:103–109 Delaunay F, Marty Y, Moal J, Samain JF (1993) The effect of monospecific algal diets on growth and fatty acid composition of Pecten maximus (L.) larvae. J Exp Mar Biol Ecol 173:163–179 Domı´nguez L, Villalba A, Fuentes J (2010) Effects of photoperiod and the duration of conditioning on gametogenesis and spawning of the mussel Mytilus galloprovincialis (Lamarck). Aquac Res 41: 807–818 Eglinton TI, Boon JJ, Minor EC, Olson RJ (1996) Microscale characterization of algal and related particulate organic matter by direct and temperature-resolved mass spectrometry. Mar Chem 52:27–54 Enright CT, Newkirk GF, Craigie JS, Castell JD (1986) Evaluation of phytoplankton as diets for juvenile Ostrea edulis (L.). J Exp Mar Biol Ecol 96:1–13 FAO (ed) (2010) The state of world fisheries and aquaculture. FAO, Rome Fernandez-Reiriz MJ, Labarta U, Albentosa M, Perez-Camacho A (1998) Effect of microalgal diets and commercial wheatgerm flours on the lipid profile of Ruditapes desussatus spat. Comp Biochem Physiol Part A 119:369–377 Figueras A (1989) Mussel culture in Spain and France. World Aquac 20(4):8–17 Figueras A (2007) Biologı´a y cultivo del mejillo´n Mytilus galloprovincialis en Galicia. Consejo Superior de Investigaciones Cientı´ficas, Madrid Folch J, Lees M, Sloane-Stanley GH (1957) A simple method for the isolation and purification of total lipids from animal tissues. J Biol Chem 226:497–509 Galley TH, Batista FM, Braithwaite R, King J, Beaumont AR (2009) Optimisation of larval culture of the mussel Mytilus edulis (L.). Aquac Int 18:315–325 Gros O, Frenkiel L, Moue¨za M (1997) Embryonic, larval, and post-larval development in the symbiotic clam Codakia orbicularis (Bivalvia: Lucinidae). Invertebr Biol 116(2):86–101 Gruffydd LD, Beaumont AR (1970) Determination of the optimum concentracion of eggs and spermatozoa for the production of normal larvae in Pecten maximus (Mollusca: Lamillibranchia). Helgol Wiss Meeres 20:486–497 Helm MM, Bourne N (2004) Hatchery culture of bivalves. A practical manual. Food and Agriculture Organization of the United Nations Publishing, Rome Izquierdo MS, Watanabe T, Takeuchi T, Arakawa T, Kitajima C (1990) Optimum EFA levels in Artemia to meet the EFA requirements of red sea bream (Pagrus major). In: Takeda M, Watanabe T (eds) The current status of fish nutrition in aquaculture, Japan Translation Center, Ltd., Tokyo, Japan, pp 221–232 Knauer J, Southgate PC (1999) A review of the nutritional requirements of bivalves and the development of alternative and artificial diets for bivalve aquaculture. Res Fish Sci 7:241–280 Lane A (1989) The effect of a microencapsulated fatty acid diet on larval production in the European flat oyster Ostrea edulis. In: de Pauw N, Jaspers E, Ackefors H, Wikins N (eds) Aquaculture—a biotechnology in progress. Bredene, Belgium, pp 657–664 Laxmilatha P, Rao GS, Patnaik P, Nageshwara Rao T, Prasad Rao M, Biswajit D (2011) Potential for the hatchery production of spat of the green mussel Perna viridis Linnaeus (1758). Aquaculture 312:88–94 Leonardos N, Lucas IAN (2000a) The nutritional value of algae grown under different culture conditions for Mytilus edulis (L.) larvae. Aquaculture 182:301–315 Leonardos N, Lucas IAN (2000b) The use of larval fatty acids as an index of growth in Mytilus edulis (L.) larvae. Aquaculture 184:155–166
123
1076
Aquacult Int (2013) 21:1065–1076
Liu W, Alabi AO, Pearce CM (2008) Fertilization and embryonic development in the basket cockle, Clinocardium nuttallii. J Shellfish Res 27(2):393–397 Loosanoff VL, Davis HC (1963) Rearing of bivalve molluscs. In: Russell FS (ed) Advances in marine biology. Academic Press, London, pp 1–136 Madrones-Ladja JA (1997) Notes on the induced spawning, embryonic and larval development of the window-pane shell, Placuna placenta (Linnaeus, 1758), in the laboratory. Aquaculture 157:137–146 Marshall R, McKinley S, Pearce CM (2010) Effects of nutrition on larval growth and survival in bivalves. Rev Aquac 2:33–55 Moue¨za M, Gros O, Frenkiel L (2006) Embyonic development and shell differentiation in Chione cancellata (Bivalvia, Veneridae): an ultrastructural analysis. Invertebr Biol 125:21–33 Muller-Feuga A, Robert R, Cahu C, Robin J, Divanach P (2003) Uses of microalgae in aquaculture. In: StrØttrup JG, MacEvoy LA (eds) Live feeds in marine aquaculture. Blackwell Publishing, Oxford, pp 253–299 Napolitano GE, Ackman RG, Ratnayake WMN (1990) Fatty acid composition of three cultured algal species (Isochrysis galbana, Chaetoceros gracilis and Chaetoceros calcitrans) used as food for bivalve larvae. J World Aquac Soc 21:122–130 Pere´z-Camacho A (1977) Experencias en cultivos de larvas de tres especies de moluscos bivalvos: Venerupis pullastra (Montagu), Venerupis decussata (Linnaeus) y Ostrea edulis (Linnaeus). Bol Inst Esp Oceanogr III 235:227–234 Pere´z-Camacho A, Labarta U, Beiras R (1995) Growth of mussels (Mytilus edulis galloprovincialis) on cultivation rafts: influence of seed source, cultivation site and phytoplankton availability. Aquaculture 138:349–362 Pettersen AK, Turchini GM, Jahangard S, Ingram BA, Sherman CDH (2010) Effects of different dietary microalgae on survival, growth, settlement and fatty acid composition of blue mussel (Mytilus galloprovencialis) larvae. Aquaculture 309:115–124 Ragg NLC, King N, Watts E, Morrish J (2010) Optimizing the delivery of the key dietary diatom Chaetoceros calcitrans to intensively cultured GreenshellTM mussel larvae, Perna canaliculus. Aquaculture 306:270–280 Rico-Villa B, Le Coz JR, Mingant C, Robert R (2006) Influence of phytoplankton diet mixtures on microalgae consumption, larval development and settlement of the Pacific oyster Crassostrea gigas (Thunberg). Aquaculture 256:377–388 Robert R, Gerard A (1999) Bivalve hatchery technology: the current situation for the Pacific oyster Crassostrea gigas and the scallop Pecten maximus in France. Aquat Living Resour 12:121–130 Robert R, Parisi G, Rodolfi L, Poli BM, Tredici MR (2001) Use of fresh and preserved Tetraselmis suecica for feeding Crassostrea gigas larvae. Aquaculture 192:333–346 Robert R, Chre´tiennot-Dinet MJ, Kaas R, Martin-Je´ze´quel V, Moal J, Le Coz JR, Nicolas JL, Bernard E, Connan JP, Le Dean L, Gourrierec G, Leroy B, Que´re´ C (2004) Ame´lioration des productions phytoplanctoniques fourrage, RI DRV/RA-2004-05, p 149 Ruiz M, Tarifen˜o E, Llanos-Rivera A, Padget C, Campos B (2008) Efecto de la temperatura en el desarrollo embrionario y larval del mejillo´n, Mytilus galloprovincialis (Lamarck, 1819). Rev Biol Mar Oceanogr 43(1):51–61 Silberfeld T, Gros O (2006) Embryonis development of the tropical bivalve Tivela mactroides (Born, 1778) (Veneridae: subfamily Meretricinae): a SEM study. Cah Biol Mar 47:243–251 Spencer BE (2002) Molluscan shellfish farming. Blackwell Publishing, Oxford Sprung M, Bayne BL (1984) Some practical aspects of fertilising the eggs of the mussel Mytilus edulis (L.). J Conseil Int Explor Mer 41:125–128 Sreenivasan PV, Rao KS, Poovannan P, Thanagavelu R (1988) Growth of larvae and spat of the green mussel Perna viridis (Linnaeus) in hatchery. Mar Fish Inf Serv Tech Ext Ser 79:23–26 Thompson PA, Guo MX, Harrison PJ (1996) Nutritional value of diets that vary in fatty acid composition for larval Pacific oysters (Crassostrea gigas). Aquaculture 143:379–391 Utting SD, Helm MM (1985) Improvement of seawater quality by physical and chemical pre-treatment in a bivalve hatchery. Aquaculture 44:125–128 Velasco LA, Barros J, Acosta E (2007) Spawning induction and early development of the Caribbean scallops Argopecten nucleus and Nodipecten nodosus. Aquaculture 266:153–165 Ve´lez A, Martı´nez R (1967) Reproduccio´n y desarrollo larval del mejillo´n comestible de Venezuela, Perna perna (Linnaeus, 1758). Bol Inst Oceanogr Univ Oriente 6(2):266–285 Widdows J (1991) Physiological ecology of mussel larvae. Aquaculture 94:147–163
123
