|
| |
|
|
|
|
||||
| Home Help Feedback Subscriptions Archive Search Table of Contents | ||||
First published online February 29, 2008
Journal of Experimental Biology 211, 969-977 (2008)
Published by The Company of Biologists 2008
doi: 10.1242/jeb.014639
Physiological and biochemical traits correlate with differences in growth rate and temperature adaptation among groups of the eastern oyster Crassostrea virginica
1 Institut de Recherche sur les Zones Côtières, 232B rue de
l'Église, Shippagan, Nouveau-Brunswick, Canada, E8S 1J2
2 Institut des Sciences de la Mer, 310 allée des Ursulines, Rimouski,
Québec, Canada, G5L 3A1
3 Institut Maurice-Lamontagne, Pêches & Océans Canada,
Mont-Joli, Québec, Canada, G5H 3Z4
* Author for correspondence (e-mail: fabrice.pernet{at}ifremer.fr)
Accepted 25 January 2008
 |
Summary |
|---|
 TOP Summary INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
Key words: genetically based intraspecific variation, growth, temperature adaptation, scope for growth, lipid remodelling, homeoviscous adaptation, mollusc, aquaculture
|
INTRODUCTION |
|---|
|
TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
).
The energy budget or `scope for growth' provides a means of integrating the
basic physiological processes into an index of energy available for growth and
reproduction. In bivalves, the scope for growth has proved to be an accurate
predictor of total production, which includes growth rate and gamete
production (for a review, see Bayne et al.,
1985) (see also Toro et al.,
1996; Labarta et al.,
1997; Smaal and Vonck,
1997; Bayne, 1999;
Bayne et al., 1999b;
Pouvreau et al., 2000;
Rueda and Smaal, 2004).
The standard metabolic rate, defined as the minimum energy requirement for
the maintenance of all essential functions within an inactive animal, is an
important component of total production
(Bayne et al., 1985).
Interestingly, the unsaturation of membrane phospholipids (the number of
double bonds per 100 fatty acid chains) is positively correlated with the
standard metabolic rate in allometric comparisons of mammals and birds
(Couture and Hulbert, 1995;
Hulbert et al., 2002b;
Hulbert et al., 2002a).
Membrane bilayers in metabolically active systems are more polyunsaturated and
less monounsaturated than those in systems that are metabolically less active
(Hulbert and Else, 1999;
Hulbert and Else, 2000). Such
polyunsaturated membranes have been proposed to result in an increased
molecular activity of membrane proteins; in this manner, the amount of
membrane and its composition can act as a pacemaker for metabolism. We have
shown that intra- and interspecific differences in basal metabolic rates in
bivalves relate to membrane unsaturation as predicted by Hulbert's theory of
membranes as metabolic pacemakers (Pernet
et al., 2006; Pernet et al.,
2007b).
The eastern oyster Crassostrea virginica is a eurythermal
suspension-feeding bivalve distributed around the east coast of North America
from the Gulf of St Lawrence to the Gulf of Mexico
(Galtsoff, 1964). In the Gulf
of St Lawrence, C. virginica is restricted to the warm shallow bays
and estuaries in the southwestern part. In these areas, water temperature
increases from below zero during the winter to ca 22–25°C
during the summer. Crassostrea virginica is of primary interest for
cold-water aquaculture because of its high commercial value. However,
variability in growth rates among individuals and sporadic juvenile
overwintering mortalities complicate the commercial exploitation of this
species (Lavoie, 1995).
Therefore, there is some interest in establishing a selective breeding
programme for cold-water performance in C. virginica, considering
that bioenergetic parameters (Hawkins et
al., 1989; Tremblay et al.,
1998; Pernet et al.,
2006), survival and growth [see Dégremont et al.
(Dégremont et al.,
2007) and references therein] are genetically correlated in many
bivalve species. However, little is known about the physiological and
biochemical responses of C. virginica to temperature.
We first hypothesized that physiological (metabolic rate and scope for growth) and biochemical (membrane lipid) traits would correlate with intraspecific growth differences in C. virginica. We predicted that fast-growing oysters allocate a smaller proportion of their aerobic capacity to maintenance requirements than do slow-growing animals. We also predicted that differences in metabolic rate among oyster groups correlate with the unsaturation index of their membrane lipids. Finally, we predicted that the scope for growth of these oysters correlates with their growth rate as the fast- and slow-growing animals differ in their capacity to exploit food.
Our second hypothesis was that genetically distinct groups of oysters
exhibit different degrees of adaptation to temperature. The underlying premise
was that physiological rates in C. virginica would show a marked
degree of temperature dependence even after a period of acclimation
(Newell et al., 1977;
Shumway and Koehn, 1982).
Furthermore, we predicted that juvenile C. virginica would counteract
the thermal effects on membrane fluidity by changing their membrane fatty
acids as predicted by the theory of homeoviscous adaptation. Briefly, the
theory of homeoviscous adaptation states that the `viscosity' of the lipid
bilayer is adjusted to offset changes imposed by temperature
(Sinensky, 1974;
Hazel, 1995;
Hayward et al., 2007). An
important component of the cold response is the increase in membrane lipid
unsaturation, and this has been linked to an enhanced resistance to cold
(Hayward et al., 2007). Given
that membrane adaptation is now viewed as a central contributor to low
temperature survival of all organisms
(Hayward et al., 2007), this
trait may be of particular interest for the cold-water aquaculture of marine
invertebrates.
|
MATERIALS AND METHODS |
|---|
|
TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
).
Hatchery oysters obtained from the Coastal Zone Research Institute (CZRI,
Shippagan, NB, Canada) were reared following a standard procedure
(Dégremont, 2003).
Briefly, 100 adults collected in February 2005 at Bouctouche (NB, Canada) were
conditioned in the CZRI hatchery. The seawater temperature was gradually
increased from –1°C to 20°C over a 15 day period and maintained
for a 35 day conditioning period. A cultured phytoplankton diet of
Isochrysis galbana, Chaetoceros gracilis and Pavlova lutheri
was added to the seawater at a total concentration of 
100 cells
µl–1. After the conditioning period, five males and 20
females were randomly selected for the production of juveniles. Spermatozoids
or oocytes were collected by stripping the gonad. Sperm from each male was
combined with the oocytes from four different females to produce 20 full-sib
families on 7th April 2005. Three-million oocytes per female were fertilized
at a ratio of 200 spermatozoids per oocyte. Additionally,
650x103 oocytes from each female were mixed together and
fertilized with a mixture of spermatozoids from each male to produce pooled
offspring (Dégremont,
2003). Larvae were reared in 115 l tanks at 21°C in 0.5
µm-filtered seawater and fed daily with the diet used for broodstock at
40 cells µl–1. Water and food were renewed three
times per week. Oyster larvae attained the pediveliger stage 22 days
post-hatching and settled on collectors that were suspended in the larval
tanks for 10 days. When hatchery-reared oysters reached at least 8 mm in shell
length (9th August), they were placed in floating bags and transferred to the
aquaculture grow-out site alongside the wild juvenile oysters. Oysters
originating from the field, the pooled group, and three full-sib families,
designated F24, F26 and F35, were harvested
on 12th October 2005 and transferred to the Station Aquicole de
Pointe-au-Père (Institut des Sciences de la mer, ISMER, QC, Canada). We
chose these five groups of oysters for their differences in growth rate, as
calculated from settlement until collection.
Oyster characterization
Growth and standard metabolic rates
Upon arrival at ISMER, 80 oysters from each group were numbered with
bee tags and placed into two 150 l aquaria (40 oysters per group per
aquarium, salinity was 28 and temperature was 20±0.8°C).
Temperature, salinity and oxygen were monitored daily with multiparameter
probes YSI 85 (Yellow Springs Incorporated, OH, USA). Oysters were fed
continuously with a mixed suspension of I. galbana, P. lutheri and
Nannochloropsis sp. at 10 cells µl–1. The shell
length of 68–74 individual oysters from each group was recorded monthly
for 9 months.
In the meantime, 50 oysters from each group were numbered with bee tags and
placed into a 150 l aquarium. After 21 days of acclimation to laboratory
conditions, the minimum oxygen consumption
(
O2min or
standard metabolic rate) was measured after having starved the oysters for 10
days. Oysters were kept individually in 50 ml metabolic chambers for 60 min
before starting the measurement. Empty shells were used as a control. Six
chambers were used simultaneously, which allowed us to measure five oysters
and one control at a time. Animals that remained closed in the chamber were
excluded from physiological analysis. Oxygen consumption for an individual
oyster was determined by sealing the chamber and measuring the reduction in
%O2 with a Strathkelvin 928 6-channel dissolved oxygen system using
a microcathode 1302 electrode (Strathkelvin Instruments, Glasgow, Scotland,
UK). Seawater was well mixed with a magnetic stirrer. The output signal was
monitored continuously on a computer until a decrease in O2 of at
least 20% was reached. After the oxygen uptake measurement, each set of
oysters was frozen at –80°C for later determination of dry mass,
which was measured after drying at 70°C for 72 h. Respiration rate is
expressed as the rate expected for a standard oyster in ml–1
g–1 dry mass h–1 by application of
allometric correction (Widdows and
Johnson, 1988).
Genetics
Upon arrival at ISMER, 50 oysters from each group were stored at
–80°C until genetic analyses. DNA extraction was carried out using
DNeasy tissue kits (Qiagen, Mississauga, ON, Canada). Genetic variation was
determined at the polymorphic microsatellite loci Cvi7,
Cvi8, Cvi 9, Cvi12, Cvi13, Cvi14
and Cvi23 (Brown et al.,
2000; Reece et al.,
2004). Amplified fragments were separated by capillary
electrophoresis using an ABI PRISM 310 automated genetic analyser (Applied
Biosystems, Foster City, CA, USA) and analysed with the Genescan and Genotyper
software packages (Applied Biosystems).
Temperature experiment
Experimental design
Upon arrival at ISMER, 243 animals from each group were equally distributed
into nine 37 l aquaria (6-Pack Arctic model, Aquabiotech, Coaticook, QC,
Canada). Each aquarium had its own filtration, aeration and water temperature
control unit. Oysters were acclimated to laboratory conditions for 5 days
prior to starting the experiment. The salinity was 28, the natural photoperiod
was followed, and the temperature was maintained at 12°C
(Fig. 1). The five groups of
oysters were maintained together in these aquaria over the entire experiment.
Animals were fed a diet of I. galbana, P. lutheri and
Nannochloropsis sp. at a total concentration of 10 cells
µl–1 twice a day. Temperature was monitored every day in
each tank throughout the experiment.
|
1.5°C day–1) to 4°C while the
temperature of three other aquaria was gradually increased (1.5°C
day–1) to 20°C. The temperature of the remaining aquaria
was maintained at 12°C as a control
(Fig. 1). All animals reached
the desired temperature by 24th October after which these temperatures were
maintained for 6 weeks. Physiological rates and lipid composition of oysters
were measured on 24th October after attaining the desired temperature (acute
response) and again on 5th December (long-term acclimation;
Fig. 1).
Physiological rates
A pool of five oysters per aquarium was used for physiological measurements
in 500 ml chambers before the oysters were killed for dry mass determination
to calculate mass-standardized physiological rates by allometric equation.
Clearance rate, defined as the volume of water cleared of suspended
particles per unit time and biomass, was quantified using a static system
(Pernet et al., 2007b).
Briefly, animals were provided with I. galbana at an initial
concentration of 10 cells µl–1 and food particles were
counted every 15 min for 60 min using an electronic particle counter (Beckman
Coulter Counter Z2, Mississauga, ON, Canada). The clearance rate (l
h–1) was then used in conjunction with the algal biomass (mg
ml–1) to estimate the amount of ingested energy, assuming
that the energy content of the diet was 23.5 J mg–1
(Widdows and Johnson,
1988).
Oxygen consumption
(O2) for an
individual animal was determined by sealing the chamber and measuring the
reduction in %O2 as described in the previous section. Respiration
was then expressed as ml O2 g–1 tissue dry mass
h–1 and then converted into energy equivalents using the
conversion factor 1 ml O2=20.33 J
(Widdows and Johnson,
1988).
Assimilation, defined as the product of ingested energy and absorption
efficiency (Widdows and Johnson,
1988), was estimated using the Conover ratio [see Conover, 1966 in
Widdows and Johnson (Widdows and Johnson,
1988)]. Food and faecal samples were filtered onto pre-combusted
pre-weighed 47 mm GFC filters that were rinsed with isotonic ammonium formate
(3.2%), dried at 80°C for 48 h, cooled to room temperature in a
desiccator, and re-weighed. Afterwards, they were combusted overnight at
450°C, cooled to room temperature in a desiccator, and finally weighed
again. This procedure provided estimates of the organic and inorganic
fractions contained in the food and faeces.
The scope for growth was estimated by subtracting the energy lost through
respiration and excretion from the energy obtained by food assimilation
(Widdows and Johnson, 1988).
As excretion represents <5% of the energy budget in most bivalves, it was
ignored.
Lipid analysis
Five oysters from each group were randomly sampled in each aquarium on 24th
October and 5th December in each temperature treatment for the determination
of lipid composition. Tissues of different oysters from the same aquarium were
pooled together to obtain ca 100 mg wet mass and stored in lipid-free
amber glass vials with Teflon-lined caps under nitrogen in 1 ml
dichloromethane at –80°C. Lipids were extracted
(Folch et al., 1957), and
classes determined as previously described
(Pernet et al., 2007b). Lipids
were separated into neutral lipids (including triglycerides and sterols) and
polar lipids (including mainly phospholipids) using column chromatography on
silica gel hydrated with 6% water (Pernet
et al., 2007b). Fatty acid methyl esters (FAME) from polar lipids
were prepared using 2% H2SO4 in methanol
(Lepage and Roy, 1984). Gas
chromatograph parameters and the procedure for FAME identification and
analysis have been described previously
(Pernet et al., 2007b).
Statistical analyses
The mean number of alleles per locus, and observed and expected
heterozygosities were calculated using the software Genetix 4.05
(Belkhir et al., 1998). The
software FSTAT (Goudet, 2001)
was used to calculate the fixation index FIS and
FST. The FIS values were tested (1000
randomizations) for significant differences from zero to assess the compliance
with the Hardy–Weinberg equilibrium. FST values were
used as an index of genetic differentiation among oyster groups. All
probability values were adjusted for multiple comparison tests using
sequential Bonferroni adjustments (Rice,
1989).
Three-way split–split plot ANOVAs were conducted to determine differences in the physiological rates and the fatty acid composition of the polar lipids in oysters, i.e. the unsaturation index and the major polyunsaturated fatty acids (PUFA), namely docosahexaenoic acid (22:6n-3) and eicosapentaenoic acid (20:5n-3), as a function of overwintering temperature, oyster origin and sampling time (Table 1). The unit of replication was the aquarium in which the temperature was applied. The main plots were temperature levels, subplots were oyster origin, and sub-subplots were sampling time. Here we used a mixed linear model, which models not only the means of our data but also their variances and covariances. The repeated option was applied to the term `time' to take into account temporal dependence (SAS 9.1.3 Help and documentation; SAS Institute Inc., Cary, NC, USA).
|
Where differences were detected, least-square means multiple comparison
tests were used to determine which means were significantly different. When
differences among groups of oysters were detected without interaction with
other factors (main effect), planned contrasts between group means were used
(Table 1). Residuals were
screened for normality using the expected normal probability plot and further
tested using the Shapiro–Wilk test. Data on scope for growth,
O2 and 20:5n-3
were log+1 transformed to achieve normality of residuals and homogeneity of
variances. Homogeneity of variance–covariance matrices was graphically
assessed. Analyses were carried out using SAS 9.1.3.
|
RESULTS |
|---|
|
TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
|
The minimum oxygen consumption or standard metabolic rate
(O2min) of
oysters collected in the field (0.34 ml O2 g–1
h–1) showed an increase of 49.5% compared with that of
hatchery-reared animals, where
O2min for the
four groups averaged 0.23 ml O2 g–1
h–1(Table 2).
It is noteworthy that the fast-growing oysters from families F24
and F26 showed lower
O2min than the
other group of oysters. Interestingly, the
O2min of oysters
exposed to laboratory conditions decreased linearly with increasing growth
rate
(O2min=–0.01xgrowth
rate+0.65; r2=0.998, N=5, P=0.001;
Table 2).
The average number of alleles was much higher in oysters from the field compared with those from the pooled group or the families (Table 2). Interestingly, oysters from F24 and F26 showed no deviation from Hardy–Weinberg equilibrium whereas the overall FIS values for oysters from other groups indicated significant heterozygote deficiencies. The FST values among the five groups of oysters varied from 0.09 to 0.30, and all pairwise comparisons were significant (P<0.05), thus indicating a high level of genetic differentiation (Table 3).
|
Temperature experiment
Physiological rates
The overall scope for growth varied as a function of oyster origin
(Fig. 2A). Fast-growing oysters
produced in the hatchery showed an average scope for growth of 46.6 J
h–1 g–1 compared with only 18.7 J
h–1 g–1 in animals from the field (contrast:
field vs hatchery, P<0.001). Although oysters originating
from the hatchery showed a similar scope for growth, it is noteworthy that
fast-growing animals from F26 showed the highest scope for growth.
More broadly, the scope for growth of oysters increased linearly with
increasing growth rate measured under laboratory conditions at 20°C for 9
months (y=2.75xgrowth rate–68.5;
r2=0.956, N=5, P=0.004).
|
|
|
60% of the total PUFA in oyster polar lipids. Overall, the level of 22:6n-3 in the fast-growing hatchery oysters (18.5% of total fatty acid) was 35.6% lower than that of oysters from the field, where it averaged 25.1% (Fig. 4A). The level of 22:6n-3 in animals collected in the field increased by 25% with a temperature decrease from 12°C to 4°C, whereas it remained constant (pooled and F26) or increased only marginally (by 8% and 13% for F24 and F35, respectively) in oysters produced in the hatchery. The level of 22:6n-3 varied as a function of time x temperature (Fig. 4B). Indeed, animals maintained at 4°C showed similar 22:6n-3 levels irrespective of sampling time whereas oysters exposed to higher temperatures showed a major decrease in 22:6n-3 following acclimation (Fig. 4B).
The effects of time, temperature and oyster origin interacted on 20:5n-3 (Fig. 4C). The level of 20:5n-3 in the fast-growing oysters produced in the hatchery (13.6%) was 54.8% higher than that of oysters originating from the field, where it averaged 8.8% before acclimation (Fig. 4C, left side). The 20:5n-3 level in oysters produced in the hatchery decreased with increasing acclimation temperature from 4°C to 20°C (Fig. 4C, right side). However, 20:5n-3 in hatchery-reared oysters maintained at 4°C remained constant irrespective of sampling time, whereas those exposed to temperatures of 12°C and 20°C showed a marked decrease in 20:5n-3 following acclimation. In contrast, 20:5n-3 in oysters from the field maintained at 4°C and 12°C remained constant irrespective of sampling time.
|
DISCUSSION |
|---|
|
TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
O2min) and in
the unsaturation index of membrane phospholipids coincides with higher growth
(as expressed by growth rate and scope for growth) in hatchery oysters.
Standard O2 and
membrane unsaturation were lower in fast-growing hatchery oysters compared
with that of slow-growing oysters from the field. Similarly, standard
O2 and
phospholipid unsaturation showed a positive correlation in comparison to wild
and selectively bred hard clams (Pernet et
al., 2006) and in comparison to mussels and oysters
(Pernet et al., 2007b). Here
we provide evidence that intraspecific differences in basal metabolic rate in
bivalves relate to membrane unsaturation as predicted by Hulbert's theory of
membranes as metabolic pacemakers (Hulbert
and Else, 1999; Hulbert and
Else, 2005).
The higher unsaturation index observed in membrane lipids of oysters from
the field was mainly due to higher 22:6n-3 levels compared with those of
fast-growing oysters from the hatchery. The importance of 22:6n-3 in
regulating an animal's metabolic rate was previously emphasized by others
(Hulbert and Else, 2005).
Mammalian phospholipids showed a statistically significant allometric decline
in the unsaturation index with increasing body size
(Hulbert et al., 2002a). This
decline in the phospholipid unsaturation index was predominantly due to the
fact that the 22:6n-3 content of tissue phospholipids decreased as the mammal
species increased in size. These results are very similar to those reported in
birds (Hulbert et al., 2002b;
Turner et al., 2006). The
functional significance of the elevated 22:6n-3 levels in the membranes of
small mammals and birds is potentially related to their high mass-specific
metabolic rate compared with larger endotherms. Because a substantial
proportion of the basal metabolism is associated with membrane-linked
processes [mitochondrial proton leak, Na+ and Ca2+
cycling together account for approximately half of standard metabolic rate
(Rolfe and Brown, 1997)], it
was suggested that membrane lipids, particularly 22:6n-3, may play a role in
determining the metabolic rate of different species via an influence
on the molecular activity of membrane-bound enzymes
(Hulbert and Else, 1999;
Hulbert and Else, 2000).
Differences in 22:6n-3 levels between oysters from the field and those from
the hatchery may reflect differences in their capacity for selective
incorporation of dietary PUFA into membrane lipids. Indeed, long-chain PUFA
such as 20:5n-3 and 22:6n-3 have been reported to be essential for sustaining
optimal growth in several bivalve species
(DeMoreno et al., 1976;
Langdon and Waldock, 1981).
Although the biosynthetic production of these PUFA is rather low or absent,
meaning that the fatty acid composition of lipids in bivalve tissues generally
reflects that of the diet, several studies have shown that the fatty acid
composition in bivalve membrane lipids is regulated by selective incorporation
or elimination of fatty acids (e.g.
Delaunay et al., 1993). It is
unlikely that differences in 22:6n-3 between groups of oysters reflect
differences in their diet as these animals were kept in the same conditions
for more than 2 months before the experiment.
Fast-growing oysters from the hatchery showed (1) a lower standard
metabolic rate and (2) a higher scope for growth due to higher food
consumption compared with slow-growing animals from the field. Similarly,
Pacific oysters Crassostrea gigas from a fast-growing line showed a
higher rate of ingestion and absorption and a lower metabolic rate at
maintenance than did those from a slow-growing line
(Bayne, 1999). A related field
experiment with the Sydney rock oyster Saccostrea glomerata (formerly
S. commercialis) showed that selection for fast-growing animals leads
to faster rates of feeding across a wide range of food concentrations
(Bayne et al., 1999b). Finally,
the superior growth in C. gigas compared with that of S.
glomerata maintained in the same conditions also coincided with a faster
rate of feeding in C. gigas
(Bayne, 1999;
Bayne, 2002). Therefore, our
results on the bioenergetics of juvenile C. virginica are in good
agreement with previously published data on other oyster species.
Finally, there was a smaller deficiency in heterozygote frequencies in
fast-growing oysters from F24 and F26 compared with that
of animals from other groups. Several studies have shown negative correlations
between heterozygote deficiency and fitness-related traits in marine bivalves,
attributed to lower metabolic requirements for heterozygous individuals
(Hawkins et al., 1989;
Tremblay et al., 1998;
Bayne et al., 1999a). In our
study, higher metabolic demands in oysters from the field, the pooled group
and the F35 associated with a higher heterozygote deficiency
probably impose a supplementary stress that results in a reduction of growth
rate in these animals.
Temperature adaptation
A perfect negative relationship was observed between acclimation
temperature and the unsaturation index of polar lipids in juvenile oysters, as
predicted by the homeoviscous adaptation theory
(Hazel, 1995). Interestingly,
changes in the unsaturation index in response to temperature were mainly due
to 20:5n-3 levels in fast-growing oysters from the hatchery, whereas
slow-growing animals changed levels of both 22:6n-3 and 20:5n-3, as previously
observed in adult C. virginica
(Pernet et al., 2007a;
Pernet et al., 2007b). This
pattern may reflect differences in the proportions and thus the availability
of 22:6n-3 and 20:5n-3 among groups of oysters (see previous section). In a
previous study, we showed that the inverse relationship between the
unsaturation index of gill phospholipids and acclimation temperature was
principally due to changes in 22:6n-3 and 20:5n-3 levels, but that the
magnitude of the response of these fatty acids varied between oysters and
mussels (Pernet et al.,
2007b). Indeed, the decrease of 20:5n-3 with a rise in temperature
was much stronger in mussels than in oysters. Although speculative, the
efficiency of homeoviscous adaptation in fast-growing oysters from the
hatchery may be higher than that of slow-growing animals from the field, since
the melting point of 20:5n-3 is 10°C lower than that of 22:6n-3.
The unsaturation index and the fatty acid composition of oysters showed
only small differences among temperature treatments on 24th October compared
with those observed on 5th December, after 6 weeks of acclimation. Therefore,
the fatty acid remodelling in the membrane lipids of juvenile oysters may be
viewed as a long-term adjustment to temperature. It is generally accepted that
changes in the fatty acid composition of membrane lipids usually occur after a
period of acclimation, varying in duration from 1 week in the case of warm
acclimation to several weeks during cold acclimation
(Hazel and Williams, 1990). A
previous study on rainbow trout showed that exposure to low temperatures led
to a gradual increase in PUFA during cold acclimation
(Hazel and Landrey, 1988).
Likewise, hard clams exposed to a lowering of environmental temperature showed
a gradual increase in the unsaturation index that was viewed as a long-term
adjustment to winter temperature (Pernet
et al., 2006). However, few studies showed that changes in fatty
acid composition could occur in the short term. For example, in oysters, the
unsaturation index of gills subjected to daily fluctuations in temperature
between 12°C and 25°C for 7 days varied in a way consistent with
membrane homeoviscous adaptation, mainly due to rapid changes in 22:6n-3 and
20:5n-3 levels (Pernet et al.,
2007a).
There was no significant temperature effect on the scope for growth of
juvenile oysters maintained at 4°C, 12°C and 20°C, reflecting the
fact that both energy expenditure (respiration) and acquisition (ingestion)
increased with temperature. This result contrasts with several published
studies on other bivalve species (for a review, see
Bayne et al., 1985) (see also
MacDonald and Thompson, 1986;
Beiras et al., 1995;
Yukihira et al., 2000;
Cusson et al., 2005). For
example, the scope for growth of juvenile oysters O. edulis that were
initially maintained at 20°C and further exposed to 14°C, 20°C or
26°C for 3 weeks increased with temperature
(Beiras et al., 1995).
Likewise, the scope for growth of mussels M. edulis acclimated at
8°C was found to be higher than that of mussels kept at –1°C
(Cusson et al., 2005).
Furthermore, considering that scope for growth is a predictor of total
production (growth rate in juveniles), our result suggest that growth rate in
oysters would be similar over temperatures ranging from 4°C to 20°C,
which is very unlikely. The lack of a temperature effect on the scope for
growth of oysters in our study may reflect an inability of the statistical
model to detect the effect of temperature due to the low number of replicate
tanks (n=3) coupled with high variation: animals acclimated at
4°C, 12°C and 20°C showed scope for growth of 24.5±15.3,
30.6±17.5 and 59.3±35.8, respectively.
For the first time, we have shown intraspecific variations in the effect of
temperature on clearance rate. Overall, the clearance rate of animals exposed
to 4°C was lower than that of oysters kept at 20°C (P=0.014).
Although very few studies have specifically examined the effect of temperature
on feeding in C. virginica, one paper reported that the pumping rate
in this species increased steadily as temperature rose from 8°C to
28°C (Loosanoff, 1958).
More recently, we showed that adult C. virginica acclimated to
overwintering temperatures (<9°C) exhibited a reduced clearance rate
compared with that of animals acclimated to spring–summer conditions at
20°C (Pernet et al.,
2007b), which agreed well with the pattern observed in
F26.
Oxygen consumption increased with temperature irrespective of sampling
time, thus suggesting that the respiration rate of these animals did not
acclimate even after 6 weeks. We know from the physiological literature that
thermal acclimation is not a universal feature of bivalve metabolism: the
ability of bivalves to alter metabolism following a temperature change is
species specific [see for example Beiras et al.
(Beiras et al., 1995) and
references therein]. More particularly, the oxygen consumption in oysters
C. virginica and Ostreas edulis remains strongly dependent
on the ambient temperature even after 3 weeks, thus suggesting that these
species have a limited ability to acclimate to temperature change
(Newell et al., 1977;
Shumway and Koehn, 1982).
Therefore, the marked thermal dependence of the respiration rates found in
this work on juvenile C. virginica is consistent with previous
studies on oyster species.
|
Acknowledgments |
|---|
|
References |
|---|
|
TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
|---|
Bayne, B. L. (1999). Physiological components of growth differences between individual oysters (Crassostrea gigas) and a comparison with Saccostrea commercialis. Physiol. Biochem. Zool. 72,705 -713.[CrossRef][Medline]
Bayne, B. L. (2002). A physiological comparison between Pacific oysters Crassostrea gigas and Sydney rock oysters Saccostrea glomerata: food, feeding and growth in a shared estuarine habitat. Mar. Ecol. Prog. Ser. 232,163 -178.[CrossRef]
Bayne, B. L., Brown, D. A., Burns, K., Dixon, D. R., Ivanovici, A., Livingstone, D. R., Lowe, D. M., Moore, M. N., Stebbing, A. R. D. and Widdows, J. (1985). The Effects of Stress and Pollution on Marine Animals. New York: Praeger.
Bayne, B., Hedgecock, D., McGolrick, D. and Ress, R. (1999a). Feeding behaviour and metabolic efficiency contribute to growth heterosis in Pacific oysters Crassostrea gigas (Thunberg). J. Exp. Mar. Biol. Ecol. 233,115 -130.[CrossRef]
Bayne, B. L., Svensson, S. and Nell, J. A. (1999b). The physiological basis for faster growth in the Sydney rock oyster, Saccostrea commercialis. Biol. Bull. 197,377 -387.[Abstract]
Beiras, R., Camacho, A. P. and Albentosa, M. (1995). Short-term and long-term alterations in the energy budget of young oyster Ostrea edulis L. in response to temperature change. J. Exp. Mar. Biol. Ecol. 186,221 -236.[CrossRef]
Belkhir, K., Borsa, P., Goudet, J., Chikhi, L. and Bonhomme, F. (1998). Genetix, logiciel sous WindowsMT pour la génétique des populations. Montpellier, France: Laboratoire génome et populations, CNRS UPR 9060, Université de Montpellier II.
Brown, B. L., Franklin, D. E., Gaffney, P. M., Hong, M., Dendanto, D. and Kornfield, I. (2000). Characterization of microsatellite loci in the eastern oyster, Crassostrea virginica.Mol. Ecol. 9,2217 -2219.[Medline]
Couture, P. and Hulbert, A. J. (1995). Membrane fatty acid composition of tissues is related to body mass of mammals. J. Membr. Biol. 148,27 -39.[Medline]
Cusson, M., Tremblay, R., Daigle, G. and Roussy, M. (2005). Modeling the depuration potential of blue mussels (Mytilus spp.) in response to thermal shock. Aquaculture 250,183 -193.[CrossRef]
Dégremont, L. (2003). Etude des bases génétiques de la mortalité estivale et des relations avec la croissance chez les juvéniles de l'huître creuse Crassostrea gigas. PhD thesis, Université de Caen, France.
Dégremont, L., Ernande, B., Bedier, E. and Boudry, P. (2007). Summer mortality of hatchery-produced Pacific oyster spat (Crassostrea gigas). I. Estimation of genetic parameters for survival and growth. Aquaculture 262,41 -53.[CrossRef]
Delaunay, F., Marty, Y., Moal, J. and Samain, J.-F. (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.[CrossRef]
DeMoreno, J. E. A., Moreno, V. J. and Brenner, K. R. (1976). Lipid metabolism of the yellow clam, Mesodesma mactroides: 2. Polyunsaturated fatty acid metabolism. Lipids 11,561 -566.[Medline]
Folch, J., Lees, M. and Sloane-Stanlez, G. H.
(1957). A simple method for the isolation and purification of
total lipids from animal tissues. J. Biol. Chem.
226,497
-509.
Galtsoff, P. S. (1964). The American oyster, Crassostrea virginica Gmelin. US Fish. Wildl. Serv. Fish. Bull. 64,480 pp.
Goudet, J. (2001). FSTAT, a program to estimate and test gene diversities and fixation indices (version 2.9.3.2). Updated from Goudet 1995. FSTAT version 1.2: a computer program to calculate F-statistics. J. Hered. 86,485 -486.
Hawkins, A. J. S., Rusin, J., Bayne, B. L. and Day, A. J. (1989). The metabolic/physiological basis of genotype dependent mortality during copper exposure in Mytilus edulis. Mar. Environ. Res. 28,253 -257.[CrossRef]
Hayward, S. A. L., Murray, P. A., Gracey, A. Y. and Cossins, A. R. (2007). Beyond the lipid hypothesis: mechanisms underlying phenotypic plasticity in inducible cold tolerance. In Molecular Aspects of the Stress Response: Chaperones, Membranes and Networks. Vol. 594 (ed. P. Csermely and L. Vigh), pp. 132-142. Berlin: Springer[CrossRef]
Hazel, J. R. (1995). Thermal adaptation in biological membranes: is homeoviscous adaptation the explanation? Annu. Rev. Physiol. 57,19 -42.[Medline]
Hazel, J. R. and Landrey, S. R. (1988). Time course of thermal adaptation in plasma membranes of trout kidney. II. Molecular species composition. Am. J. Physiol. 255,R628 -R634.[Medline]
Hazel, J. R. and Williams, E. E. (1990). The role of alterations in membrane lipid composition in enabling physiological adaptation of organisms to their physical environment. Prog. Lipid Res. 29,167 -227.[CrossRef][Medline]
Hulbert, A. J. and Else, P. L. (1999). Membranes as possible pacemakers of metabolism. J. Theor. Biol. 199,257 -274.[CrossRef][Medline]
Hulbert, A. J. and Else, P. L. (2000). Mechanisms underlying the cost of living in animals. Annu. Rev. Physiol. 62,207 -235.[CrossRef][Medline]
Hulbert, A. J. and Else, P. L. (2005).
Membranes and the setting of energy demand. J. Exp.
Biol. 208,1593
-1599.
Hulbert, A. J., Rana, T. and Couture, P. (2002a). The acyl composition of mammalian phospholipids: an allometric analysis. Comp. Biochem. Physiol. 132B,515 -527.[CrossRef][Medline]
Hulbert, A. J., Faulks, S., Buttemer, W. A. and Else, P. L.
(2002b). Acyl composition of muscle membranes varies with body
size in birds. J. Exp. Biol.
205,3561
-3569.
Labarta, U., Fernandez-Reiriz, M. J. and Babarro, J. M. F. (1997). Differences in physiological energetics between intertidal and raft cultivated mussels Mytilus galloprovincialis.Mar. Ecol. Prog. Ser. 152,167 -173.[CrossRef]
Langdon, C. J. and Waldock, M. J. (1981). The effect of algal and artificial diets on the growth and fatty acid composition of Crassostrea gigas spat. J. Mar. Biol. Assoc. U. K. 61,431 -448.
Lavoie, R. E. (1995). Culture of the American oyster, Crassostrea virginica. In Cold-water Aquaculture in Atlantic Canada (ed. A. D. Boghen), pp.191 -226. Moncton, NB, Canada: The Canadian Institute for Research on Regional Development.
Lepage, G. and Roy, C. C. (1984). Improved recovery of fatty acid through direct transesterification without prior extraction or purification. J. Lipid Res. 25,1391 -1396.[Abstract]
Loosanoff, V. L. (1958). Some aspects of
behavior of oysters at different temperatures. Biol.
Bull. 114,57
-70.
MacDonald, B. A. and Thompson, R. J. (1986). Influence of temperature and food availability on the ecological energetics of the giant scallop Placopecten magellanicus III. Physiological ecology, the gametogenic cycle and scope for growth. Mar. Biol. 93,37 -48.[CrossRef]
Newell, R. C., Johson, L. G. and Kofoed, L. H. (1977). Adjustment of the components of energy balance in response to temperature change in Ostrea edulis.Oecologia 30,97 -110.[CrossRef]
Pernet, F., Tremblay, R., Gionet, C. and Landry, T.
(2006). Lipid remodeling in wild and selectively bred hard clams
at low temperatures in relation to genetic and physiological parameters.
J. Exp. Biol. 209,4663
-4675.
Pernet, F., Gauthier-Clerc, S. and Mayrand, E. (2007a). Change in lipid composition in Eastern oyster (Crassostrea virginica Gmelin) exposed to constant or fluctuating temperature regimes. Comp. Biochem. Physiol. 147B,557 -565.[CrossRef][Medline]
Pernet, F., Tremblay, R., Comeau, L. and Guderley, H.
(2007b). Temperature adaptation in two bivalve species from
different thermal habitat: energetic and remodeling of membrane lipids.
J. Exp. Biol. 210,2999
-3014.
Pouvreau, S., Bacher, C. and Heral, M. (2000). Ecophysiological model of growth and reproduction of the black pearl oyster, Pinctada margaritifera: potential applications for pearl farming in French Polynesia. Aquaculture 186,117 -144.[CrossRef]
Reece, K. S., Ribeiro, W. L., Gaffney, P. M., Carnegie, R. B.
and Allen, S. K. (2004). Microsatellite marker development
and analysis in the eastern oyster (Crassostrea virginica):
confirmation of null alleles and non-Mendelian segregation ratios.
J. Hered. 95,346
-352.
Rice, W. R. (1989). Analysing tables of statistical tests. Evolution 43,223 -225.[CrossRef]
Rolfe, D. F. S. and Brown, G. C. (1997).
Cellular energy utilization and molecular origin of standard metabolic rate in
mammals. Physiol. Rev.
77,731
-758.
Rueda, J. L. and Smaal, A. C. (2004). Variation of the physiological energetics of the bivalve Spisula subtruncata (da Costa, 1778) within an annual cycle. J. Exp. Mar. Biol. Ecol. 301,141 -157.[CrossRef]
Shumway, S. E. and Koehn, R. K. (1982). Oxygen consumption in the American oyster Crassostrea virginica. Mar. Ecol. Prog. Ser. 9,59 -68.[CrossRef]
Sinensky, M. (1974). Homeoviscous adaptation-a
homeostatic process that regulates the viscosity of membrane lipids in
Escherichia coli. Proc. Natl. Acad. Sci. USA
71,522
-525.
Smaal, A. C. and Vonck, A. (1997). Seasonal variation in C, N and P budgets and tissue composition of the mussel Mytilus edulis. Mar. Ecol. Prog. Ser. 153,167 -179.[CrossRef]
Toro, J. E., Vergara, A. M. and Gallegillos, R. (1996). Multiple-locus heterozygosity, physiology and growth at two different stages in the life cycle of the Chilean oyster Ostrea chilensis. Mar. Ecol. Prog. Ser. 134,151 -158.[CrossRef]
Tremblay, R., Myrand, B., Sevigny, J.-M., Blier, P. and Guderley, H. (1998). Bioenergetic and genetic parameters in relation to susceptibility of blue mussels, Mytilus edulis (L.) to summer mortality. J. Exp. Mar. Biol. Ecol. 221, 27-58.[CrossRef]
Turner, N., Haga, K. L., Else, P. L. and Hulbert, A. J. (2006). Scaling of Na+,K+-ATPase molecular activity and membrane fatty acid composition in mammalian and avian hearts. Physiol. Biochem. Zool. 79,522 -533.[CrossRef][Medline]
Widdows, J. and Johnson, D. (1988). Physiological energetics of Mytilus edulis: scope for growth. Mar. Ecol. Prog. Ser. 46,113 -121.[CrossRef]
Yukihira, H., Lucas, J. S. and Klumpp, D. W. (2000). Comparative effects of temperature on suspension feeding and energy budgets of the pearl oysters Pinctada margaritifera and P. maxima. Mar. Ecol. Prog. Ser. 195,179 -188.[CrossRef]
CiteULike 
Complore 
Connotea 
Del.icio.us 
Digg 
Reddit 
Technorati 
Twitter What's this?
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
