© 2005 International Council for the Exploration of the Sea
Growth, behaviour, and digestive enzyme activity in larval Atlantic cod (Gadus morhua) in relation to rotifer lipid
Ocean Sciences Centre, Memorial University of Newfoundland St. John's, NL, Canada A1C 5S7
*Correspondence to C. C. Parrish: tel: +1 709 737 3225; fax: +1 709 737 3220. e-mail: cparrish{at}mun.ca.
Atlantic cod (Gadus morhua) show great potential for aquaculture, but much is unknown about their digestive capacity and efficiency. An integrated experiment was performed on cod larvae to investigate the variability in digestive development in response to the quantity of lipid in the rotifer enrichment. Survival, growth, behaviour, and digestive enzyme data from hatching to metamorphosis [0–450 dd (degree-days)] were measured. Four treatments were used in triplicate: high lipid rotifer enrichment (HLRE), low lipid rotifer enrichment (LLRE), green water, and unfed. Swimming activity and attacks (captures + misses) on prey were higher in the HLRE group at 100 dd than in other treatments, and this difference increased thereafter. There was no difference in digestive enzyme activity between the unfed and greened treatments, while the LLRE larvae had lower activity levels than larvae fed HLRE by 100–150 dd for all enzymes assayed. The larvae in the unfed and green water treatments did not survive past 100 dd. All the LLRE cod had died by 250 dd. Results suggest that a higher quantity of lipid in the rotifer enrichment will not only promote better growth and survival in Atlantic cod larvae but appears to provide more energy, allowing larvae to capture more live prey.
Keywords: attacks, cod digestive capacity, growth, rotifer enrichment, survival, swimming activity
Received 13 June 2004; accepted 21 November 2005.
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Introduction |
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Atlantic cod (Gadus morhua) is being studied in several countries as a species that might lead to commercial aquaculture production in the near future. Cod have good biological potential for mass culture because they are highly fecund, spawn readily in captivity, and are capable of growth rates similar to other cultured species, even at low temperatures (Howell, 1984; Finn et al., 2002). To achieve good growth, two factors influence the way food is used as a metabolic fuel by a rapidly growing larval fish: the nutritional content of the food and the efficiency with which the larva can digest and metabolize nutrients (Jobling, 1994).
Studies of the growth of larval marine fish have stressed the importance of successful initiation of foraging behaviour during the critical first-feeding period (Hunter, 1981; van der Meeren and Naess, 1993; Hunt von Herbing and Gallager, 2000). It is essential to larval growth that basic energetic and growth demands are met, which entails prey being captured in sufficient numbers to meet essential nutritional requirements (Puvanendran and Brown, 1999). In the mass rearing of cod larvae, the rotifer Brachionus plicatilis is used as a first-feeding prey, but it alone is not a sufficiently complete diet to meet metabolic demands (Rainuzzo et al., 1989).
For Atlantic cod, optimizing live feed enrichments first requires an understanding of the digestibility and retention of three major nutrient classes: carbohydrates, proteins, and lipids. Carbohydrates are not a major component of the diet of larval cod, and as such, cod larvae lack digestive carbohydrase activity during the first two months of life (Perez-Casanova, 2003). Thus, research is now focused predominantly on the importance of proteins and lipids.
During the previous 20 years, studies on live food enrichments for larval teleosts have focused on the quantity and quality of lipids in their formulas. It has been reported that high quantities of lipids in the first-feeding diet will improve larval development (Rainuzzo et al., 1997). Lipids are important as they form the basis of cellular membranes and are vital to proper neural development, but they also transport lipid-soluble compounds such as vitamins and are structural precursors for hormones (Jobling, 1994). Further, increasing the overall energy of the diet is considered beneficial to high feeding efficiency, protein sparing, and to reducing phosphorous and nitrogen losses (Kaushik and Olivia-Teles, 1985; Cahu et al., 2000). Diet enrichments focus on providing larvae with essential lipids for development (Rainuzzo et al., 1997; Cahu et al., 2000), while at the same time not providing excessive amounts of lipids that may become deposited in the liver and increase production costs (Lie et al., 1986; Grant et al., 1998; Morias et al., 2001).
Recent studies have shown trends in the development of digestive enzyme activities over time in larvae such as sea bass (Dicentrarchus labrax) (Infante and Cahu, 1994), Atlantic cod, and haddock (Melanogrammus aeglefinus) (Perez-Casanova, 2003). However, knowledge of the ability of larvae to adapt and vary their digestive development in response to dietary differences has not been widely investigated (Hoehne-Reitan et al., 2001) and is lacking for larval cod. Furthermore, enzymatic activity has not been studied and analysed concurrently with growth and behavioural data in this respect. This experiment addresses the question, does the level of lipid in rotifer enrichment affect the survival, growth, and behavioural and biochemical responses of Atlantic cod larvae.
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Material and methods |
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Larviculture and diets
Atlantic cod eggs were obtained from broodstock located at Ocean Sciences Centre (Logy Bay, Newfoundland, Canada), disinfected by immersion in PerosanTM for 60 s, thoroughly rinsed in seawater for several minutes, and incubated in 500-l tanks at 6°C until
70% had hatched [80 degree-days (dd)], which was taken as day 0 of the experiment. Cod were transferred and acclimated to 3000-l experimental tanks, stocked with 50 larvae l–1 with constant aeration at 8°C. An initial water flow of 2 l min–1 and low light intensity were gradually increased to 8 l min–1 and 2000 lux, respectively, over the first two weeks of the experiment. Temperature ranged from 8°C to 10°C over the course of the experiment in the 3000-l tanks. Rotifers (B. plicatilis) were used as live prey for the duration of this experiment. Rotifers were reared in 3000-l tanks, starting with 450 l freshwater and 450 l seawater at 23–28°C, with constant gentle aeration. Initial stocking density was less than 700 ml–1. Rotifers were cultured on baker's yeast, Saccharomyces cerevisiae, and the culture Selco. Each day, a population and egg count were performed on a number per millilitre basis, then 420 l of water (315 l filtered seawater and 105 l freshwater) was added to the tank. This was repeated daily for 6 days, after which the rotifers were removed from the tank, washed, concentrated, and placed in a 300-l conical tank for enrichment.
Two diets were used in triplicate: a high lipid rotifer enrichment (HLRE), with 21% dry weight (dw) TAG (triacylglycerol), and a low lipid rotifer enrichment (LLRE), with 5% dw TAG. Single tanks were "greened" with Isochrysis algae only, or were unfed, without replicates for ethical reasons. Enrichments were added to the rotifer tanks twice in a 12-h period at 09:00 and 15:00. The enriched rotifers were harvested at 09:00, rinsed, and put into 10 l seawater in 20-l buckets and kept in the same room as the experimental tanks. Air stones were added to the buckets to provide constant aeration. Ten litres of seawater was added to the buckets to further dilute the rotifers and reduce their temperature. This helped prevent rotifers from settling out of the tank before larvae had an opportunity to feed. Aliquots of 10 ml were taken at several locations throughout the tank three times daily and counted to determine prey density. Rotifers were added to the tanks as necessary to sustain an optimal prey density of 4000 l–1 (Puvanendran and Brown, 1999). There were typically <500 rotifers l–1 in the tanks when quantified just before the 09:00 feeding.
Enrichment analyses
Rotifer samples were taken at the 09:00 feeding time from the feed containers, not the larval tanks. The rotifers were enriched twice as per manufacturer's directions at 21:00 and 15:00, so the sampling for lipids and fatty acids was done 6 h after the second enrichment period. During the first week of the experiment, triplicate samples of rotifers (unenriched rotifers to serve as a baseline for comparison, and the LLRE and HLRE rotifers) were taken for dry weight and lipid analyses. Rotifer samples were placed directly in chloroform and stored at –20°C until processed. Lipids were extracted in chloroform/methanol according to Parrish (1998), using a modified Folch procedure (Folch et al., 1957). Lipid classes and fatty acids were determined by thin-layer chromatography with flame ionization detection (TLC/FID) with an Iatroscan (Parrish, 1987). Extracts were spotted on silica gel-coated Chromarods, and a three-stage development system was used to separate lipid classes.
Survival
Survival at the end of the experiment was determined first by allowing each treatment tank to drain slowly until the volume of water had decreased from 3000 l to 500 l. Then the fish remaining in the tank were counted in one-fourth of the tank area, and this number was multiplied by four to obtain the total cod surviving in each treatment tank. Three estimates of survival were determined for each treatment tank. Estimates among replicate treatment tanks, which were not found to be significantly different from each other at the end of a given treatment (p > 0.05), were combined to determine a mean number of cod survival.
Growth and behaviour
Growth data for standard length and myotome height were taken every 50 dd (7 days) with a stereomicroscope and a calibrated eyepiece micrometer. Condition factor and length-specific growth rate were calculated as per Jobling (1994):
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Behavioural data were taken every 50 dd (7 days) using a 2-min focal animal technique, where an individual larva is observed for 2 min in the larval feeding tank. Observations were conducted within 10 min of the first morning feeding. "Swimming activity" was defined as the percentage time a larva moved through the water column by movements of the caudal body area, and "attacks" were defined as the frequency (number per time) of captures (bites and ingestions) and misses (failed captures) of prey (Puvanendran and Brown, 2002). A Psion computer observation was used to record and compile data. Ten larvae were observed per tank per treatment.
Digestive enzyme activity data
Triplicate pooled samples of larvae were taken for biochemical determination of digestive enzyme activity at 0, 20, 50, 100, 150, 200, 250, 300, 350, 400, and 450 dd post-hatch. Three hundred larvae were collected at hatch to obtain the minimum 0.5-ml tissue necessary for each assay, and the number of fish collected over time decreased as the fish grew. After repeated visual verification, larvae were collected between 09:30 and 10:00 to ensure empty guts and to reduce the impact of prey enzymes. A small aquarium net was used to remove larvae from the tanks, and fish were transferred to a 1-l cup with 0.07 mg l–1 of MS-222 in seawater. A plastic 1-ml pipette was used to remove larvae from the cup by gently sucking them up into the transparent neck of the pipette; following this, larvae were counted. Then larvae were released onto a framed mesh of 500-µm netting. Saltwater and MS-222 were allowed to drain off the fish, which were then rinsed with freshwater and also drained. The larvae were finally placed in 1.5-ml Eppendorf tubes and stored at –80°C until they were processed and analysed.
Processing the tissues for analysis required defrosting the samples on ice (4°C), diluting with four parts (wt/vol) 150 mM NaCl, then using an automated tissue grinder (Polytron homogenizer by Brinkman Instruments with a standard generator and saw teeth) to create whole body homogenates. These homogenates were transferred to 1.5-ml Eppendorf tubes and centrifuged for 10 min at 12 000 g at 4°C. The supernatants were aliquoted in 0.5-ml Eppendorf tubes and stored at –80°C until assayed for total protein concentration, general proteases, trypsin-like alkaline proteases, pepsin-like acidic proteases, general lipases, and alkaline phosphatase. Assays were conducted in triplicate for each replicate sample and were performed at room temperature by diluting with two volumes of the respective assay buffer.
Total protein concentration of the homogenate was assayed with the BioRad Protein Assay (Bradford, 1976). Bovine gamma globulin was used to produce a standard curve from which concentrations were derived. Then these total protein concentrations were used to standardize the subsequent enzyme activity results to larval size as the fish grew over the course of the experiment.
General protease activity was measured using the azocasein hydrolysis assay (Ross et al., 2000). Samples were incubated in 2.5 volumes of 5.0 mg ml–1 azocasein dissolved in 100 mM ammonium bicarbonate (pH 7.8). Samples were shaken constantly for 20 h at 30°C, after which the reaction was stopped by adding 0.3 volumes of 20% TCA (trichloroacetic acid). These mixtures were centrifuged for 10 min at 17 000 g, and 100 µl of supernatant was added to 100 µl 0.5 M NaOH in a microplate. Optical density was measured at 450 nm. Enzyme-specific activities are reported as units of activity per milligramme total protein. (One unit represents the amount of enzyme that will increase the absorbance of the sample 0.001 optical density units over 20 h.)
Trypsin-like alkaline protease activity was assayed, based on the methods in Gawlicka et al. (2000). The substrate was prepared by diluting 4.4 mg BAPNA in 50 µl DMSO, then adding 5 ml of a 100 mM ammonium bicarbonate buffer (pH 7.8) to yield a 2 mM BAPNA solution. Cod samples (25 µl) were incubated with 50 µl of the 2 mM BAPNA substrate and 25 µl buffer for 5 min at room temperature (25°C) in a microplate. The blank contained 50 µl buffer and 50 µl substrate. Optical density was then measured at 450 nm in 30-s intervals over 30 min to get an initial rate of reaction, which was then used to calculate the enzymatic activity of the sample. Enzyme activities are reported as units of activity per milligramme protein. (One unit represents 1 µmol p-nitroaniline liberated during 1 min of hydrolysis.)
Pepsin-like acidic protease activity was assayed according to methods in Anson (1938). The substrate was prepared from a mixture of 0.2 g of 2% haemoglobin stock solution in 10 ml distilled water, mixing vigorously, and filtering through a glass wool filter. Eight millilitres of this solution then had 2 ml HCl added to adjust the pH to 2.0. Cod samples (50 µl) were incubated for 10 min at room temperature (22°C) with 250 µl of the final 1.6% haemoglobin substrate in a microplate. The reaction was stopped by adding 500 µl of 5% (wt/vol) TCA, and then centrifuged at 3600 g for 6 min. The blank contained 250 µl substrate and 500 µl 5% TCA. Optical density was measured at 280 nm on the supernatant in quartz cuvettes. Enzyme activities are reported as units of activity per milligramme total protein. (One unit represents 1 µmol tyrosine liberated during 1 min of hydrolysis.)
General lipase activity was assayed according to methods in Gawlicka et al. (2000). A detergent solution was prepared by adding 1.0 ml of 10% Triton X-100 to 9.0 ml 100 mM ammonium bicarbonate buffer (pH 7.8). The substrate was a 10 mM solution of p-nitrophenyl myristate in 100% ethanol. A microplate containing wells with 25 µl buffer, 25 µl sample, and 50 µl detergent solution was incubated at room temperature (22°C) for 15 min, then 4 µl of room temperature cod samples was added. Blanks contained 50 µl buffer, 50 µl detergent solution, and 4 µl substrate. Optical density was measured at 405 nm in 30-s intervals over 30 min to get an initial rate of reaction, which was then used to calculate the enzymatic activity of the sample. Enzyme activities are reported as units of activity per milligramme protein. (One unit represents 1 µmol p-nitrophenol liberated during 1 min of hydrolysis.)
Alkaline phosphatase activity was assayed according to methods in Gawlicka et al. (2000). A 100 mM ammonium carbonate–magnesium chloride buffer was made by adding 0.79 g ammonium bicarbonate to 0.020 g magnesium bicarbonate and diluting in 100 ml distilled water. The substrate was a 20 mM solution of p-nitrophenyl phosphate disodium prepared by diluting 74.22 mg p-nitrophenyl phosphate disodium in 10 ml of the buffer solution. A microplate containing wells with 55 µl buffer and 25 µl sample was incubated at room temperature (22°C) for 5 min, then 20 µl of room temperature substrate was added. Blanks contained 80 µl buffer and 20 µl substrate. Optical density was measured at 405 nm in 30-s intervals over 30 min to obtain an initial rate of reaction, which was then used to calculate the enzymatic activity of the sample. Enzyme activities are reported as units of activity per milligramme protein. (One unit represents 1 µmol p-nitrophenol liberated during 1 min of hydrolysis.)
Statistical analyses
Prior to analysis, survival, growth, behaviour, and enzyme data were tested for assumptions of normality and homogeneity of variance before an ANOVA (analysis of variance) was employed. Distribution of the data, plots of residuals, and predicted values were examined. There was no significant tank effect (F2,8 < 4.07, p > 0.05 for all treatments), and data from treatments were pooled. Statistical analyses were performed with Minitab statistical software (version 13). One-way ANOVAs and Tukey's HSD multiple range tests were performed to detect differences between the means of treatments using a significance level of 
= 0.05. The data were normal and homogeneous, so transformation or randomization was not required before statistical analyses were employed.
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Results |
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Enrichment analysis
The HLRE contained the highest quantity of overall lipids when standardized to dry weight of the sample, and the LLRE rotifers were not significantly different from the unenriched rotifers (Table 1). The proportions of triacylglycerols and free fatty acids were higher in the HLRE, while the LLRE had proportionately more acetone mobile polar lipids. The relative amounts of ethyl esters, sterols, and phospholipids were similar for rotifers in both enrichments.
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Three fatty acids were evaluated: arachidonic acid (AA, 20:4
-6), eicosapentaenoic acid (EPA, 20:5-3), and docosahexaenoic acid (DHA, 22:6-3), as well as the sum of the 3 and 6 fatty acids. In all cases, the LLRE did not differ from the unenriched baseline rotifers, but the HLRE rotifers were significantly higher than both the LLRE and unenriched rotifers (Table 1). The DHA:EPA ratio for the LLRE was 0.29, while for HLRE it was 3.71. The 3:6 ratio for the LLRE was 0.92, while the HLRE ratio was 2.43; however, AA concentrations were the same in both unenriched and HLRE rotifers.
Survival and growth
Larvae in the green water and unfed treatments died 100 dd after hatching. All three tanks containing cod larvae fed the LLRE died by 350 dd. A total of 4613 ± 1358 (mean ± standard error) larvae survived in the HLRE treatment at 450 dd (3.1% survival).
Both myotome and standard length for the unfed and green water treatments decreased relative to their size at hatch. Myotome height was significantly higher for larvae fed HLRE by 150 dd (F1,59 = 28.06, p < 0.001), and standard length was significantly greater by 200 dd (F1,59 = 14.54, p < 0.001) compared with that of larvae fed LLRE. Myotome height and standard length for LLRE larvae reached a plateau between 100 and 300 dd. HLRE larvae, however, continued to grow over the course of the experiment (Figure 1A and B).
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Condition factor showed a similar trend to that of length. HLRE-fed larvae had significantly higher condition factors than LLRE larvae by 200 dd (F1,59 = 39.68, p < 0.001). The HLRE larvae continually increased condition factor over the course of the experiment, while the LLRE larvae displayed a slow overall decline in condition factor, and both green water and unfed treatments decreased condition until death at 100 dd (Figure 2A).
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Length-specific growth rates were negative after the initial sampling date for greened and unfed larvae, and both LLRE and HLRE larvae slowed their growth rates over the course of the experiment (Figure 2B). However, the rate of growth slowed significantly faster in the LLRE larvae by 200 dd (F1,59 = 20.80, p < 0.001) compared with the HLRE larvae (Figure 2B).
Behaviour
The proportion of time larvae spent swimming was not significantly different between the greened and unfed treatments, where both treatments declined to 100 dd. Swimming activity for larvae fed HLRE was fairly constant. By the end of the experiment, HLRE larvae spent 38.19% of their time swimming, while larvae fed LLRE significantly reduced swimming activity after 100 dd (F1,5 = 12.49, p = 0.024) and the activity declined until death (Figure 3A).
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With the exception of 150 dd, attacks on rotifers were observed to be significantly lower in LLRE-fed larvae after 100 dd (F1,5 = 11.96, p = 0.026), while HLRE larvae increased their attacks up to 200 dd, after which the predation rate stabilized (Figure 3B). Attacks on prey were not measured in the green water and unfed tanks, since no rotifers were introduced to the tanks.
Digestive enzyme activity
The activity of digestive enzymes differed among treatments. The unfed and greened treatments had their highest enzyme activity levels at hatching, followed by a constant decline in activity for all enzymes measured (proteases, trypsin, pepsin, lipases, and alkaline phosphatase), as well as a decline in overall protein concentration of the larvae (Figures 4–9
). Comparing the activity of digestive enzymes between cod fed rotifers enriched with either HLRE or LLRE, a similar pattern was observed. Since the larvae fed LLRE did not survive past 250 dd, there are no data for that treatment beyond that time.
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What is interesting, however, is the point at which the enzyme activity levels start to differ significantly. The HLRE larvae had significantly higher values for general proteases (F1,8 = 36.39, p < 0.001; Figure 5) and alkaline phosphatase (F1,8 = 27.18, p < 0.001; Figure 9) by 100 dd, and for overall protein content (F1,8 = 30.58, p < 0.001; Figure 4), trypsin (F1,8 = 44.75, p < 0.001; Figure 6), pepsin (F1,8 = 66.12, p < 0.001; Figure 7), and lipase activity (F1,8 = 102.58, p < 0.001; Figure 8) by 150 dd. By 250 dd of age, larvae fed LLRE had reached their lowest activity levels of the experiment, similar to levels found in green water and unfed treatments before their death at 100 dd.
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Discussion |
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Cod in the HLRE treatment showed the best survival, whereas the LLRE fish only survived until 250 dd, and the unfed and greened treatments did not survive past 100 dd. The final survival rate of 3.1% in the HLRE tanks may seem low until considering the large numbers of larvae needed for the biochemical assays. At least 0.5 ml of wet tissue was required for each assay replicate. At hatch, this resulted in 300–350 larvae collected in triplicate from each tank. The numbers of fish necessary to make up the minimum volume of tissue decreased as individual larval mass increased. As such, tank populations were not only affected by diet but also by the disturbance of removing individuals for sampling. However, equal disturbances were performed in all tanks on a given sample day, so the effect of sampling was similar among the treatments. The numbers of larvae removed from tanks during the course of the experiment were not deducted from this final survival rate. When the large sampling factor is accounted for by subtracting the number of larvae removed, the final survival rate increases to 36.2%.
Performance data indicate that HLRE is a better live prey diet than LLRE prey, since all replicates of LLRE-fed cod died by 250–300 dd. However, mortality in the low lipid treatment was preceded by a predictable pattern of decreased growth, condition, and foraging activity. Growth and condition of the larvae were significantly higher in the high lipid treatment by 150–200 dd. Larval swimming activity and prey ingestion rates were also significantly higher in the HLRE treatment by 100 dd.
Activity data are a good first indication of larval performance (Skiftesvik, 1992) since fish that are not receiving energy to meet metabolic demands have less energy to expend for activities like swimming or prey capture, even when the prey swim slowly and in predictable patterns and are easily caught by saltatory predators like cod larvae (Buskey et al., 1993; Hunt von Herbing and Gallager, 2000). The HLRE larvae grew and consistently increased their overall growth as well as swimming activity and attack rate. Cod larvae in the LLRE treatment, however, showed characteristics similar to those of starved larvae, such as decreased foraging activity and increased buoyancy (Laurence, 1978; Kjorsvik et al., 1991; Skiftesvik, 1992). Tissue degradation was found in other species of marine fish larvae under similar conditions of starvation (Yin and Blaxter, 1987). This would suggest that even though ample prey were available, the larvae in this experiment did not possess the energy required to ingest prey given that the energy derived from previous foraging activity was not sufficient to support metabolic demands of growth.
Ellertsen et al. (1980) reported that cod larvae, at the onset of exogenous feeding, had a successful predation rate of 32–62%, which increased to over 90% by the end of yolk-sac absorption. As such, even though a high activity level would increase the odds of ingesting live prey, low activity levels in cod larvae may be a strategy to conserve energy and cope with starvation, or at least delay the onset of irreversible starvation.
In addition to decreased foraging activity, the digestive efficiency of cod can be inefficient under poor feeding conditions. If sufficient nutrition is not available to a larva, impaired development of the fins results, as well as the inability of the fish to increase its overall size (and manoeuvrability), and rapid degradation of the digestive tissues occurs (Ellertsen et al., 1980; Yin and Blaxter, 1987; Kjorsvik et al., 1991). Concurrent with tissue degradation is a reduction in the ability of the gut to process food for energy. With respect to this experiment, even if larvae were still capable of prey capture in the early stages of starvation, reduced activity and incomplete digestion resulting from gut tissue degradation likely contributed to the onset of starvation. The time to irreversible starvation of cod has been reported as 70 dd when no food was available (Laurence, 1978). In this experiment, when nutritionally deficient food was available significant differences in survival, growth, and behaviour were apparent at 100 dd. Given that the live feed was the same in both rotifer treatments, the data suggest that the quantity or quality of lipid in the enrichment, and not the live prey itself, led to decreased survival, growth, swimming activity, and ingestion rates. Qualitatively, the high lipid diet was similar to the low lipid rotifer treatments in terms of ethyl esters, triacylglycerols, sterols, phospholipids, and AA proportions or concentrations.
A positive correlation has been found for DHA:EPA ratios and larval growth in yellowtail flounder (Limanda ferruginea) (Copeman et al., 2002) and gilthead sea bream (Sparus aurata) (Rodriguez et al., 1997). Mirroring the lipid composition of marine fish eggs has been suggested as a starting point for determining nutritional requirements of newly hatched larvae. A typical DHA:EPA ratio of 2:1 has been found in several marine larval species and suggested as adequate for larval feeding (Sargent et al., 1999). In the current experiment, the LLRE had a DHA:EPA ratio of 0.3, which is equivalent to that of unenriched rotifers. The HLRE, however, had a significantly higher ratio of 3.7. In light of the growth, lipid, and behavioural data in this experiment, the LLRE diet was deficient in vital lipids. In herring (Clupea harengus), diets deficient in DHA change the fatty acid composition of neural tissues and decrease foraging efficiency (Bell et al., 1995). Atlantic cod possess high levels of DHA in both eye and brain tissues (Bell and Dick, 1991). Since cod are visual feeders, inadequate amounts of DHA may inhibit their ability to forage successfully. This deficiency of DHA in the eye tissue of cod larvae leading to visual impairment could be a major factor explaining the marked decline in larval cod foraging activity by 100 dd. This decreased foraging activity was followed by slower growth measures and digestive enzyme activities compared with the HLRE larvae by 150 dd and death by 250 dd.
Digestive enzyme activity reflected the ability of the larvae to digest available food items for metabolic energy and was similar to general patterns found in other larval species, such as Senegal sole (Solea senegalensis) (Ribiero et al., 1999). It is difficult, however, to make meaningful comparisons between studies that use vastly different rearing temperatures, diets, sampling methods, and assays to determine digestive enzyme activity. The biochemical assays of the current experiment used similar methods to, and results concur with, those of Perez-Casanova (2003). In both cases, trypsin- and pepsin-like enzymes were all present and active at hatching. The previous study, however, examined activity with one feeding regime. The results of the current experiment show similar overall trends in enzyme activity for both types of proteases, but the activities of lipase and alkaline phosphatase were higher in cod larvae fed HLRE in this experiment than cod larvae in the Perez-Casanova (2003) study. The overall patterns of digestive enzyme activity in the two experiments are similar, but differences in enrichments elicited distinct biochemical responses. Additionally, Perez-Casanova (2003) compared the contribution of rotifer digestive enzymes to that of the whole body homogenates of Atlantic cod larvae. Rotifer enzymes contributed 16.4% to the total general lipase activity, 10.8% to trypsin-like enzyme activity, 3.5% to general protease activity, and 0% to pepsin-like activity. As such, the amount of digestive enzymes in the rotifer live food was minimal and did not significantly impact the results of the enzyme assays.
Enzyme activity levels obtained from the trypsin and pepsin assays are not specific for these enzymes, as they detect trypsin-like alkaline proteases and pepsin-like acid proteases. Furthermore, the assays were performed on whole body homogenates, which further complicates interpretation as interference may result from proteases and protease inhibitors in tissues other than from the digestive system. One would not expect to see pepsin activity in early larval development since the larvae do not have a functional stomach until metamorphosis. The resultant activity of pepsin-like enzymes in this experiment is most likely the result of other acidic proteases like aspartic proteases in the pepsin family of digestive enzymes.
The activities of general lipases and alkaline phosphatases in the HLRE treatment were higher than in the study by Perez-Casanova (2003) for larvae beyond 250 dd. There are two possible explanations for this result: (i) the increase in activity is the result of a decrease in feeding activity as the larvae approach metamorphosis, initiating an increase in the need to hydrolyze stored lipids for energy (Martinez et al., 1999), or (ii) feeding cod larvae a diet with high lipids for energy induces the digestive system to increase its lipid-digesting capabilities. Data from the behavioural portion of this experiment do not support the first option, but instead support the theory that increases in enzyme activity are in fact the result of the influence of food. Since alkaline phosphatase has been generally accepted as a marker of intensity of nutritional absorption in the intestine of larvae teleosts (Segner et al., 1989), the results of this experiment will have implications in assessing the digestive capacity of fish if this increase in enzyme activity is the result of diet and not feeding activity.
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Acknowledgements |
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We thank the Aquaculture Research Development Facility (ARDF) staff for their valuable help with live-food production and larval rearing. This work was funded by AquaNet – Canada's Network of Centres of Excellence for aquaculture research.
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Footnotes |
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Deceased 4 September 2005. 
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References |
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