© 2004 by ICES/CIEM International Council for the Exploration of the Sea/Conseil International pour l'Exploration de la Mer
Discarding Norway lobster (Nephrops norvegicus L.) through low salinity layers – mortality and damage seen in simulation experiments
a Department of Biology, University of Leicester Leicester LE17RH, UK
b Institute of Marine Research PO Box 4, 453 21 Lysekil, Sweden
*Correspondence to R. R. Harris; tel: +44 116 2523341; fax: +44 116 2523330. e-mail: rrh{at}leicester.ac.uk; Mats.Ulmestrand{at}fiskeriverket.se.
1tel: +46 523 187 00; fax: +46 523 139 77.
The Kattegat/Skagerrak Nephrops fishery is unusual in that animals normally live in high salinities (33–34 psu) but are raised through a low salinity surface layer and, if discarded, descend back through it to the sea bed. In other open-sea Nephrops fisheries, such low salinity exposure is rare. Physiologically the species is described as being marine stenohaline, i.e. intolerant of reduced salinities, and a lower salinity limit of 29–30 for its distribution has been suggested. Using CTD data from the Anholt E hydrographic station, near to known Nephrops grounds, a knowledge of hauling times (including washing the cod-end with surface water) and of the sinking rate of Nephrops, we simulated conditions experienced by the catch in this fishery. We also included a period of emersion (air exposure) on deck during sorting or reshooting the trawl. Blood electrolytes, body mass changes and simple behavioural responses were examined before and after the simulation over a 5-day period. "Discarded" Nephrops experienced significant haemodilution and gained mass rapidly. Animals showed slow rates of "tail-flipping", or absence of responses to stimulation, in the period immediately following return to salinity 33 seawater, although many showed recovery later. Delayed effects included abdominal stiffness, swelling and further mortalities (25–42% overall). Controls (exposed to 33 psu seawater only) showed good survival and vigorous responses, even with a period of emersion. The effects of salinity exposure, which are additional to the stresses of being trawled, on the fitness of discarded animals were evaluated. Our results suggest that Nephrops discard mortality in this fishery is significantly higher than past estimates and is due to the stress of this brief exposure to low salinity surface water.
Keywords: damage, discards, fishery, Kattegat/Skagerrak, low salinity stress, mortality, Nephrops norvegicus
Received 3 March 2003; accepted 18 August 2003.
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1 Introduction |
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 Top 1 Introduction 2 Materials and methods3 Choice of low...4 Results5 DiscussionReferences |
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Commercially valuable Norway lobster Nephrops norvegicus (L.) fisheries occur around the northeastern Atlantic continental shelf and in the Mediterranean. The species lives in burrows in suitable mud sediments at depths ranging from 15 to 800 m (Farmer, 1975; Chapman and Howard, 1988), and is caught either by otter trawling or by creel-fishing (traps). One of the features of Nephrops fisheries is the high percentage of discards in the catch; either non-target species or Nephrops which are below a size restriction imposed upon the fishery for management reasons. The minimum landing size (MLS) of 40 mm carapace length in the Skagerrak/Kattegat area is considerably higher than the MLS of 25 mm in the North Sea area. This implies that more than half of the average Nephrops catch in the former area is below MLS and is discarded back into the sea. The survival of target species discards is important for ensuring the maintenance of the Nephrops stock and estimates of discard mortality are used in stock assessments. The ICES Nephrops Assessment Working group have used a figure of 75% discard mortality (ICES, 1997), although many measurements made on the species have been slightly lower (in the range 60–69%) (Symonds and Simpson, 1971; Redant and Polet, 1994; Wileman et al., 1999). These mortality estimates are based on experiments in somewhat protected conditions (sea bed cages or tanks on deck), and take no account of predation by either fish or seabirds following discarding from the deck and sinking back to the sea bed. Discarding often takes place away from Nephrops grounds increasing discard susceptibility to predation at sites where refuges are scarce. Also discards in poor condition have been shown to be less successful in competing for food and shelter compared to apparently undamaged conspecifics (Evans et al., 1994).
Many factors are responsible for the poor condition and mortality of discards. Physical damage due to abrasion and compression within the cod-end can result in major injuries which vary according to catch size, composition, trawl duration and speed (Bergmann et al., 1998; Wileman et al., 1999). Treatment on-deck can also contribute to discard mortality. Long-periods of emersion (aerial exposure) during sorting of the catch can cause internal hypoxia, with resulting lactic acid and ammonia build-up, body fluid dehydration and concentration (Spicer et al., 1990; Wileman et al., 1999). These physiological changes may have profound effects on survival and recovery may be slow (Wileman et al., 1999; Bergmann and Moore, 2001). Elevated on-deck temperatures and high light intensities have also been shown to have damaging effects (Zainal et al., 1992; Chapman et al., 2000).
The Kattegat/Skagerrak Nephrops fishery is unusual in that animals normally live at high salinities (33–34) but are raised, either in creels or trawls, through a low salinity surface layer and, if discarded, are allowed to descend back through it to the sea bed. In other Nephrops fishery areas (Scotland West Coast, Irish Sea, Iberian continental shelf slope, Mediterranean) such low salinity exposure is rare. CTD data show that a halocline occurs throughout the year at 10–15 m in the Southern Kattegat with minimum surface water salinities varying between 15 and 25. Further north in the southern Skagerrak, surface salinities are slightly higher (17.5–33 depending on season). During the summer and autumn, surface salinities rise above the minimum as Baltic water outflow rates decrease. This is the case also in winter when freshwater is locked in the Baltic as ice. Bottom water salinities are consistently in the range 33–34 in most parts.
Thus, in addition to the stresses of trawling, exposure of discards to low salinities in these fisheries could be a cause of stress and mortality. Reports on the salinity tolerance of Nephrops are few. Early studies of its ecology suggested a lower salinity limit of 29–30 for its distribution (Poulsen, 1946; Farmer, 1975). This is well above the surface salinities described in the area. It has been described as a marine stenohaline species normally not found (or surviving) in salinities very different from full-strength seawater (Mantel and Farmer, 1983; Schoffeniels and Dandrifosse, 1994).
The aim of the present study was to assess the effects of low salinity exposure on discard mortality and physiological stress by means of a simple simulation of being lifted through and sinking back through a low salinity layer. Treatments were based upon typical conditions and times of exposure derived from field data obtained in the Kattegat/Skagerrak area. Periods of emersion were based upon a knowledge of commercial operations and two emersion temperatures were also included. Observations of behaviour, tail-flip escape responses and measurement of important haemolymph (blood) parameters were carried out to assess the survival, fitness and recovery of Nephrops trawled and discarded under these conditions.
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2 Materials and methods |
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Top1 Introduction2 Materials and methods3 Choice of low...4 Results5 DiscussionReferences |
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Small adult Nephrops, generally falling within the size range of animals which would be discarded in this Nephrops trawl or creel fishery (<40 mm CL) (mean wet mass = 35.94±0.90 g; n=72), were caught by baited creels in the Skagerrak near the mouth of Gulmarsfjorden, West Sweden (58°15'N 11°26'E) at about 40–55 m depth in the first week in April 1999 (Figure 1). Field conditions in that period were: bottom salinity = 33.13±0.09 psu; bottom temperature = 5.14±0.05°C (mean ± SE; n=15); air temperature at noon = 13.4±1.2°C (data courtesy of Kristineberg Marine Biological Station, Fiskebäckskil, Sweden). Animals were transferred to flowing seawater (33 psu; 5°C) in holding tanks at Kristineberg. They were held separately in opaque polyethylene tubes (25 cm length x 5.5 cm diameter), each drilled along its length with at least five 1 cm diameter holes to allow water circulation. The tube ends were closed by pieces of 17 mm mesh nylon netting held in place with rubber bands for easy removal when sampling. All tubes were numbered to ensure rapid identification of individuals.
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Animals were kept in these tubes throughout the pre-experimental acclimation period (2 weeks), and during all experimental transfers to different salinity conditions. Survival within the tubes in the holding tanks was 100%, and in control conditions (see below) mortality was also minimal. Individuals could be examined daily with the least disturbance by gently lifting one end of the tube and examining the animal's head end. In holding conditions, animals were fed with the flesh of one Mytilus placed within each tube on alternate days. Each experiment commenced with a period of 46.67 h in 33 psu seawater when animals were unfed. Groups of six Nephrops within tubes were placed in glass aquaria (60x30x30 cm) connected to the cooled, circulating seawater supply in a temperature-controlled room (contained volume of seawater = 27 l; salinity = 33, temperature = 5°C). Animals were removed at intervals for weighing and haemolymph sampling. At this time the condition of each animal was also assessed visually and scored using the following scale: animals in Condition 1 showed active, strong tail-flipping, Condition 2 animals showed slow tail-flipping, but with some appendage movement, Condition 3 animals showed complete abdominal immobility but with some reflexes in appendages when touched, and Condition 4 animals showed no reflexes, nor any scaphognathite activity, and appeared moribund or dead.
Experimental animals were exposed to a laboratory simulation of being trawled and discarded on local Nephrops grounds. Thus the changes in salinity and temperature, including a period of emersion (air exposure), that animals would undergo when being brought to the surface in a trawl through a low salinity surface layer were recreated. We also included in the protocol a period of time to simulate discarding (as undersized) from the deck of a trawler and sinking through a low salinity surface layer, eventually returning to the sea bed.
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3 Choice of low salinity exposure times |
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Top1 Introduction2 Materials and methods3 Choice of low...4 Results5 DiscussionReferences |
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A knowledge of CTD data at the Anholt E sampling station (56°40'N 12°07'E) in the Kattegat was used to design the simulation experiments (Figure 1) (courtesy of the Swedish Meteorological and Hydrological Institute). This station is near to known Nephrops grounds. Figure 2 shows salinity/depth profiles measured during the months of April to July 1998 at this station. Based on discussions with local fishermen, we calculated an average haul rate of 7.2 m min–1 (haul time differed between large and small trawlers and with the depth, which is between 50 and 160 m). From a knowledge of the extent of the low salinity layer, we calculated that trawled Nephrops would spend a maximum of 6 min within it when being brought to the surface. This would include a period of washing the cod-end with surface water, at the ship's side, to remove sediment. Subsequently animals would spend an average of 90 min on deck during sorting during which time they may be exposed to higher temperatures, particularly in summer. A small catch would be sorted more rapidly but sorting might be delayed while shooting the next trawl. During this time some low salinity water would probably be held within the gill chambers of Nephrops. Discarded animals would sink back through the surface layers to the high salinity water below. To estimate this time interval spent in the surface layers, the sinking rates of Nephrops were measured in groups of 10 animals which had been both submerged and emersed prior to measurement. This was carried out in a large glass cylinder (0.5 m diameter and 1.5 m in height). Animals were released at the surface and their rate of fall (m min–1) calculated from the time taken to sink to depths of 1 and 1.5 m. Sinking experiments were carried out in 15 and 33 salinity seawater, and at two temperatures (5 and 15°C). No significant differences were observed between mean sinking rates of animals in either of the two salinities, or at the two different temperatures, (mean ± SE in 15 at 5°C = 8.88±0.27; at 15°C = 8.59±0.22 m min–1; n=30 in each case). Emersion produced a minor, but statistically non-significant, increase in sinking rate (9.67±0.82 and 8.82±0.21 m min–1). On the basis of these observations, a time of 98 min (6 min + 90 min + 2 min) was selected for low salinity exposure periods in the following treatments.
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Groups of 12 animals were exposed to the following:
- Controls in seawater (CSW; Table 1) were held in the conditions referred to above in flowing 33 psu seawater at 5.0 (±1.0)°C for 120 h. Body masses were determined and haemolymph samples taken at 24, 48, 72, 96 h.
- Simulation with emersion at 5°C animals (EM/5) were held in 33 psu seawater at 5.0±1.0°C for 46.67 h. The medium was then replaced with 27 l of 15 psu seawater (diluted by addition of distilled water to full-strength seawater; 33) at 5.0°C for 6 min. Subsequently, animals were moved in their tubes into air (emersion) and placed in a plastic fish basket at the same temperature for 90 min. They were then replaced in 15 psu seawater for 2 min, before being returned to 33 psu seawater for the remainder of the experiment. Animals were sampled at the times described in (a).
- SUB/5 animals (at 5°C with no emersion) were held in 33 psu seawater at 5.0°C for 46.67 h. The medium was replaced with 15 psu seawater. Animals were held at 5.0°C for 98 min (1.63 h) continuously in this medium and then returned to 33 psu seawater. There was no period of emersion. Animals were sampled at the same intervals as in (a).
- EM/15 animals (simulation with emersion at 15°C) were held in 33 psu seawater at 5.0°C for 46.67 h. The medium was then replaced with 15 psu seawater at 15 (±1)°C for 6 min. Animals were then placed, within their tubes, into a plastic fish basket and held emersed at 15°C for 90 min. Following this they were returned to 15 psu seawater at the same temperature for 2 min, and then replaced in 33 psu seawater at 5.0°C for the remainder of the experiment.
- SUB/15 animals were treated similarly except that submersion in 15 psu was for 98 min continuously, with no period of emersion in air; the whole period in 15 salinity was at a temperature of 15°C.
- A control-in-air group (CEM) was also included in which animals were held in 33 psu seawater at 5.0°C for 46.67 h then emersed in a fish basket for 90 min at 15°C, then returned to 33 psu seawater at 5.0°C.
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Table 1 summarizes the conditions and duration of exposures for the five animal treatment and control groups. For all groups, visual assessments of animal condition were made at the sampling times and, additionally, at 120 h when animals were tested for tail-flip escape swimming performance by a method adapted from Newland et al. (1988) and Field (1992). Animals were transferred individually into an opaque plastic aquarium (0.9x0.5x0.5 m), containing well-aerated seawater. They were removed from their individual tubes with a minimum of disturbance by gently encouraging the animals to walk out. They were then stimulated to produce tail-flip escape swimming by frequent tapping of the rostrum and anterior margins of the cephalothorax, with a hand-held plastic rod, until no further response was obtained. The total number of tail-flips (flexion and extension of abdominal segments) elicited and the duration of these responses were recorded. Exhaustion was defined as a lack of response to three successive final taps.
Animals were wet-weighed (±0.1 g) after superficial water was removed and drained from the gill chambers by gentle shaking with the animal held in a head-down position. Successive weighings of control animals showed that this procedure gave consistent results, with an error of 0.59% for daily weighing of the same individuals (n=12) over a 5-day period. Haemolymph samples (
0.5 ml) were removed in air using 1 ml disposable plastic hypodermic syringes from the base of a walking leg, transferred immediately to Eppendorf tubes and shaken vigorously for 2 min. Sampling was generally completed in <30 s to minimize emersion and the stress of handling. Samples were then centrifuged (x5000g) to separate-off clotting proteins and the supernatant used for all analyses. Samples were stored frozen (–20°C) until required.
Haemolymph samples were also taken from Nephrops in the field on board the R/V "Argos" (Institute of Marine Research, Sweden) immediately after being trawled, and after a period of 90 min emersion on deck before being discarded. Lobsters were trawled from about 40 m depth at a station 56°47'68N, 11°51'11E in the Kattegat on 15 September 1999 (Figure 1). CTD profiles at this station showed surface and bottom salinities averaging 17.5 and 33.5, respectively, and temperatures of 17.5 and 11.0°C, with a surface low salinity layer extending down to 15 m depth. Samples were removed by 1 ml disposable hypodermic syringe and frozen (–15°C). They were later transported to the University of Leicester for analysis.
For all haemolymph and seawater samples, [Na+] was determined by flame photometry (Jenway PFP7) after appropriate dilution, [Cl–] by Corning 925 titrator (20 µl samples in both cases), and osmotic concentration by freezing-point depression (Roebling micro-osmometer). [K+] was measured by atomic absorption spectrophotometry (Varian AA6), following dilution (50 µl in 10 ml deionized water) of both standards (5–50 mmol l–1) and haemolymph. Salinities (psu) were measured using a WTW Microprocessor Conductivity Meter LF196.
All data are expressed as means ± 1 SE. The number of experimental animals is given in parenthesis. Differences between means were tested for significance by one-way analysis of variance following an Fmax test for homogeneity of variances (Sokal and Rohlf, 1995). Significantly different means (P<0.05) were identified using Tukey's pairwise comparison. Differences in condition and total numbers of tail-flips were tested using Kruskal–Wallis tests.
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4 Results |
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Top1 Introduction2 Materials and methods3 Choice of low...4 Results5 DiscussionReferences |
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4.1 Animal mortality and condition following low salinity exposure
Control animals maintained throughout the test period in 33 salinity seawater showed good survival (CSW = 92%) even following a 90 min period of emersion (air exposure) at 15°C (CEM = 100%) (Figure 3). However, in all treatment groups experiencing a 98 min period of exposure to 15 salinity, either with emersion (EM/5 and EM/15) or continuous submergence (SUB/5 and SUB/15), major disturbances in behaviour and changes in the lobsters' responses to stimulation were seen immediately on return to 33 seawater (at 48 h). Animals also showed significant mortality. All treatments showed significant differences in condition compared with controls (P<0.001; Kruskal–Wallis test). Figure 3 shows the number of animals in each group exhibiting each of these conditions at each sampling interval following exposure. Animals showed reduced flexure of the abdomen which sometimes had a swollen appearance with a complete absence of tail-flipping (Condition 3). Others showed slow rates of tail-flipping (Condition 2) compared with controls which consistently showed vigorous, rapid rates of tail-flipping (Condition 1). At 48 h there were no significant differences in condition between any of the simulation groups (Kruskal–Wallis test) with 58–83% of individuals showing Condition 3. At 72 h (24 h after return to 33 seawater), EM/5 and SUB/5 animals (both at 5°C) showed some recovery with a rise in the proportion of animals showing Conditions 1 and 2 but some mortality also occurred. In EM/15 and SUB/15 animals a similar recovery was seen, but 17 and 25% of these groups, respectively, died and were removed. Mortalities continued during the course of the experiment, finally attaining 42 and 25% in EM/5 and SUB/5 animals, respectively, and 25 and 33% in EM/15 and SUB/15 animals. In the latter two groups a significantly larger proportion of individuals were in Conditions 2 and 3 and fewer in Condition 1 at 120 h than was the case for other simulations. A number of individuals developed a whitish, opaque abdomen, in addition to showing an absence of tail-flip responses in Condition 3. Controls animals (CSW and CEM) showed good survival and the great majority showed vigorous tail-flipping (Condition 1) throughout the experimental period.
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4.2 Changes in body mass following low salinity exposure
Control animals (CSW and CEM) held in 33 salinity seawater throughout the experimental period (except for weighing and haemolymph sampling) showed no significant changes in body mass. Handling and withdrawal of
0.5 ml haemolymph at 24 h intervals appeared to have no important effects on body mass under these conditions (Figure 4a). In contrast, animals experiencing low salinity (15) conditions, either with a period of emersion or by continuous submersion (EM/5 and SUB/5, respectively), showed highly significant gains in mass (expressed as % original wet weight). In EM/5 animals the mean mass increased to 102.5±0.49% when measured immediately after removal (48 h) (P<0.001), while in SUB/5 animals a similar significant gain was seen (P<0.01). Similarly EM/15 and SUB/15 (low salinity at 15°C) showed significant gains in mean mass following low salinity exposure. When comparing all treatments, it was found that the extents of the mass gain immediately following exposure were not significantly different. Emersed control animals (CEM) showed a non-significant reduction in mean body mass after emersion (Figure 4a and b).
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After low salinity exposure all animals were returned to 33 salinity seawater until the termination of the experiment at 120 h. During this period, animals in the majority of treatments regained mass. The exceptions were: EM/5 animals which showed a significantly lower mean mass than initial at 72 h (24 h after exposure) (97.8±0.47% initial), and SUB/15 where mass remained at 98.4±0.39% initial at 72 h. At the final measurement (96 h), surviving animals of all treatment showed no significant differences in their masses compared to initial levels.
4.3 Changes in haemolymph solutes following low salinity exposure
Control animals held in 33 salinity throughout the experimental period maintained haemolymph osmotic concentrations slightly, but significantly, hypoosmotic to the surrounding seawater (lower than) (haemolymph = 949.5±2.0 mOsm kg–1, n=24; 33 seawater = 958.1±3.49 mOsm kg–1, n=8) (P<0.05). Transfer to 15 psu seawater (457.9±3.18 mOsm kg–1) of EM/5 and SUB/5 animals was followed by a rapid significant reduction in haemolymph osmotic concentration (P<0.001) (Figure 5a). Animals were not sampled immediately prior to the low salinity exposure period to minimize handling stress, but assuming that haemolymph osmotic concentrations before transfer were similar to those recorded at 24 h for each group, this would represent a mean fall of 140 and 182 mOsm kg–1, or 14.9 and 18.8% of the original osmotic concentration, respectively. EM/5 animals were emersed for 90 min with only 8 min of low salinity submergence, yet a large and significant reduction in osmotic concentration occurred in this group compared to CSW animals sampled at the same intervals (P<0.001). The mean haemolymph osmotic concentration recorded immediately after exposure was not significantly different from SUB/5 animals (P>0.05), in which continuous exposure to 15 psu seawater was maintained throughout 98 min. Following a return to 33, haemolymph osmotic concentrations in both EM/5 and SUB/5 animals increased to levels not significantly different from those of controls within 24 h. These levels were also maintained for the remainder of the experiment.
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Similarly, EM/15 and SUB/15 animals showed highly significant decreases in haemolymph osmotic concentration following low salinity exposure (at 15°C) with average decreases observed being 138 and 164 mOsm kg–1, respectively (P<0.001) (Figure 5b). There was no significant difference in haemolymph osmotic concentration between the two groups during recovery in 33 psu seawater. Thus, at both temperatures, being emersed or submerged during the low salinity exposure period had no significant effect on the degree of haemodilution seen. Following return to 33 psu seawater, haemolymph osmotic concentrations in EM/15 and SUB/15 animals also recovered to near original levels. However, in emersed control animals (CEM) there was a highly significant haemodilution 48 h following the period of emersion even though the salinity was maintained at 33 throughout (P<0.001).
Control animals held in 33 salinity seawater throughout the experimental period maintained a mean haemolymph [Na+] concentration of 451.0±4.5 mmol l–1 (n=24), slightly hyperionic to seawater (higher concentration than) (33=429.0±4.9; 15=220.4±7.4 mmol l–1 [Na+]; n=8). Transfer to 15 psu seawater of EM/5 and SUB/5 animals was followed by large decreases in haemolymph [Na+] (Figure 5c and d) (P<0.001). Again, surprisingly, there was no significant difference in haemolymph [Na+] between animals subjected to continuous exposure to 15 salinity seawater and those briefly submerged, with an intervening period of emersion. The largest decrease in [Na+] was seen in EM/15 animals (=295.07±3.17 mmol l–1 at the end of the low salinity exposure period; a fall of 156 mmol l–1). CEM animals showed a significant rise in haemolymph [Na+] (P<0.01). On return to 33 salinity, survivors in all experimental groups showed increases in haemolymph [Na+] to original levels, except EM/15 which had significantly lower levels than other groups (P<0.01).
Haemolymph chloride showed a similar reduction and return to original control levels following simulation of fishing and discarding through a low salinity layer (Figure 5e and f). However, there were differences, compared with the total solute and [Na+] changes, in that the largest decrease in [Cl–] was seen in SUB/5 animals which had significantly reduced haemolymph [Cl–] compared with EM/5 animals (emersed for the majority of the period of their low salinity exposure). EM/15 and SUB/15 animals showed lesser falls in haemolymph [Cl–]. In emersed control animals (CEM) haemolymph [Cl–] rose significantly above the initial levels and was higher than CSW animals (P<0.01).
Haemolymph [K+] was maintained a level below that of the surrounding 33 psu seawater in CSW animals (mean = 6.5±0.2 mmol l–1; medium = 11.5±1.1 mmol l–1 [K+]; n=8). Transfer of EM/5 and SUB/5 animals to low salinity conditions had no immediate effect on haemolymph [K+] (not significantly different from initial controls) but there was a transient rise 24 h later. Thus at 72 h haemolymph [K+] was significantly higher in EM/5 and SUB/5 animals than controls (P<0.01) but these were not significantly different from each other. A similar picture emerged in EM/15 and SUB/15 animals which showed even greater increases in haemolymph [K+] (P<0.01). In the case of SUB/15 animals, a sustained elevation of haemolymph [K+] was observed at 96 h which was about 3.5 fold higher than initial levels (Figure 5g and h). It appears that following low salinity exposure at a higher temperature, haemolymph [K+] rose to levels greater than those seen in similar salinity conditions at 5°C.
4.4 The effect of low salinity on tail-flip swimming performance
Control animals (CSW) tested at the end of the experimental period were able to perform a mean of 126.7±5.1 (11) tail-flips to exhaustion (as defined in Materials and methods). Following transfer to low salinity conditions there was a significant reduction in performance in surviving EM/5 animals (emersed at 5°C), in spite of the 96 h recovery period in salinity 33 (P<0.001) (Table 2). However, the mean number of tail-flips seen in SUB/5 animals at this time (108.0±28.76, n=9) was not significantly lower than that seen in the controls (CSW). In some cases, surviving animals showed no tail-flips at all. These were predominantly females. A similar picture emerged when EM/15 and SUB/15 animals were tested and compared with emersed controls (CEM). CEM animals showed strong tail-flipping abilities (mean 133.4±5.9 (12)) after experiencing a 90 min period of emersion whereas significant reductions in EM/15 and SUB/15 animals were seen (P<0.05). No significant differences in performance were seen when comparing EM/5 and SUB/5, nor when EM/15 and SUB/15 were similarly tested. The duration over which tail-flipping occurred showed no major change following low salinity exposure in EM/5, SUB/5 and EM/15 animals. Thus the number of tail-flips was reduced but these could be elicited over a period of time similar to that of the responses seen in CSW and CEM animals. The exception was the SUB/15 group in which a significant reduction in duration compared to controls was seen (P<0.05).
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4.5 Haemolymph parameters measured in fished Nephrops
Osmotic concentration and electrolyte concentrations of the haemolymph sampled from recently landed Nephrops, and a separate group of captured animals which had been held on deck emersed in air for 90 min before sampling (air temperature = 16.4°C), are compared with those of control animals (submerged in
33 seawater) in Table 3. Osmotic concentrations of haemolymph taken immediately after capture (807.3±11.6 mOsm kg–1 (n=20)) were significantly lower than the levels shown by control animals in the simulation study (CSW and CEM), suggesting some loss of haemolymph solute during trawling and/or hauling (P<0.001). The ambient deep-water salinity was recorded simultaneously as 33.5. Similarly, low haemolymph [Cl–] was seen in these animals (P<0.001). However, haemolymph [Na+] was not significantly reduced and [K+] was significantly higher (P<0.001). Comparisons were also made between immediate post-capture solute concentrations and those present after 90 min emersion on deck. Although all showed some increase, only mean osmotic concentration and [K+] were significantly elevated (P<0.05).
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5 Discussion |
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Top1 Introduction2 Materials and methods3 Choice of low...4 Results5 DiscussionReferences |
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No attempt was made in this study to simulate the full stresses of capture of Nephrops by trawling. These have been studied in other fisheries in the absence of low salinity exposure and, in addition to physical damage (including body fluid losses), have been shown to result in major physiological and metabolic disturbances, arising from the high levels of swimming activity within the trawl (Wileman et al., 1999). The effects of low salinity exposure in trawled animals would be additional to these disturbances, probably resulting in stresses involving osmotic and ionic perturbations.
The relatively brief low salinity exposure period used here resulted in significant mortalities immediately on removal from the discard simulation conditions and return to full-strength seawater. These were followed by additional mortalities during the following days so that, by the end of the experiments, between 42 and 25% of individuals had died depending on treatment. Furthermore, animals exposed to our simulation showed swelling of the abdomen and major disturbances in behaviour immediately post-exposure (e.g. reduced tail-flipping on stimulation and slow movements). Similar symptoms of low salinity exposure have been reported in other stenohaline marine crustaceans (Dall, 1974; Shaner et al., 1985). In some studies, mortality was described as being due to "bursting" where bulk fluid losses through the exoskeleton occurred. Body mass increases have also been reported and, in some species, these were restored to the original with time (Schwabe, 1933; Cornell, 1980). We found mass gains of a similar order in Nephrops suggesting a large and rapid uptake of water. It has been suggested that rigid-bodied decapods are susceptible to bursting due to a build-up of fluid pressure within the body during sudden low salinity exposure (Davenport, 1985). Whether or not this increase in turgor is a cause of the reduced rate and frequency of abdominal flexures seen in "tail-flipping" remains to be investigated. It is clear from our observations that the characteristic escape response of the species will be weak during and following descent through a low salinity layer, and that this will result in an increased susceptibility to predation by fish and avian predators (Evans et al., 1994). This will have further impact on discard survival in addition to the direct mortality effects seen in our experiments.
Measurements of the osmotic and ionic composition of the haemolymph (blood) of Nephrops have been made (Robertson, 1949, 1960, 1961), and of the permeability characteristics of its gill cuticle. The latter has been shown to be relatively "leaky" to Na+, Cl–, HCO3– and NH4+ (Lignon and Gendner, 1988). "Leaky" stenohaline marine animals lose blood electrolytes and gain water rapidly when transferred to low salinities as shown in early studies on the spider crab Maia squinado (Schwabe, 1933; Kirschner, 1991). Generally there is a tendency to come to a rapid osmotic equilibrium with the dilute seawater with decreases in blood osmotic and inorganic ion concentrations.
No studies of the tolerance to, or osmotic regulation of Nephrops in, low salinities have been found in the literature therefore it was necessary to carry out some preliminary studies to obtain basic data. Nephrops attained osmotic equilibrium in 28 (with 100% survival) and 25 salinity seawater (50% survival), but in a salinity of 21, animals showed 100% mortality before haemolymph concentrations had fallen to external medium levels. In our discard simulation experiments, the concentrations of major solutes (including osmotic concentrations) in the haemolymph of Nephrops fell rapidly, with the exception of K+ which showed some rises in the later phases. A comparison of these concentrations measured immediately following low salinity exposure showed that animals had not attained isosmoticity or an isoionic state (= [ion] between haemolymph and external medium) with the low salinity simulation medium. Furthermore, solute concentrations in animals which appeared moribund were higher by about 100 mOsm kg–1 (difference in [Na+] = 52; [Cl–] = 63 mmol l–1) than the levels recorded from animals in 25 psu seawater in the preliminary experiments. This suggests that during the simulation period, haemolymph concentrations had not fallen to some critical low level since those animals which were maintained long-term in 25 salinity showed lower values (mean = 768 mOsm kg–1). It is probable that the "rate" rather than the "extent" of this decrease in solute concentration is the factor determining survival following low salinity exposure. In this respect the possibility of some damage to excitable cell functioning (muscle or nerve) should be examined. Treherne (1980) reported a marked reduction in the amplitude of action potentials in leg nerves of the stenohaline crab M. squinado subjected to hypoosmotic seawater. This effect, caused by osmotic swelling of the axons, was reversible, providing the osmotic concentration of the fluid bathing the axons did not fall below the lethal limit for the species. An increase in K+ permeability of the nerve membrane caused a hyperpolarization which was responsible for reducing action potentials. An increased outward leakage of K+ from swelling cells may account for the rise in haemolymph [K+] seen in Nephrops following low salinity exposure, particularly at 15°C. Elevation of haemolymph [K+] has been reported in discarded animals not subjected to low salinity exposure, suggesting tissue damage during capture (Wileman et al., 1999). Whether or not the symptoms of diminished tail-flip responses and slow limb movements were due to an osmotic disturbance of excitable cell functioning remains to be determined.
Significantly, water content gain (as measured by mass) and haemolymph solute losses by Nephrops exposed to low salinities occurred at similar rates whether or not there was a significant period of air emersion. This suggests that a pool of low salinity water is available around the exchanging surfaces of the animal (probably the gills) which is sufficient in volume to allow significant osmotic uptake of water into the body, causing mass gain and, probably, a simultaneous outward leakage of solutes (mainly ions). Animals submerged continuously for 98 min in low salinity medium showed changes which were similar in magnitude to emersed Nephrops. Thus animals fished through low salinity layers and emersed in air on deck will have low salinity water trapped extracorporeally (probably in their gill chambers). This will continue to exchange with the haemolymph, and probably other body fluid compartments, and will result in net water gain and solute depletion. In emersed control animals, maintained in 33 salinity seawater before and after the simulation, mass changes were small (slight reduction but non-significant) while haemolymph solute concentrations showed little immediate change. It would be interesting to test, in the field, whether or not immersing the catch in full-strength seawater prior to sorting and discarding would reduce the extent of body fluid and solute concentration changes by replacing extracorporeal water with a similar volume of high salinity seawater. Following return to full-strength seawater body water content and haemolymph solute concentrations of low salinity exposed animals were restored to initial levels and maintained thus for the remainder of the experimental period. In addition to gains in body mass due to water uptake, it is probable that the dramatic dilution of the extracellular fluid (haemolymph) will cause significant osmotic stress to many of the body tissues in Nephrops resulting in osmotic flow of water into, and swelling of, cells.
Generally, rates of mortality and osmotic and ionic changes showed no significant differences in animals subjected to the discard simulation at the two temperatures 5 and 15°C, apart from the steep rise in haemolymph [K+] seen in the latter group. However, at the termination of the experiments (120 h), the EM/15 and SUB/15 groups were in poorer condition compared to EM/5 and SUB/5 animals suggesting a damaging effect of the warmer temperature used. Nephrops is generally regarded as intolerant of warm temperatures, and capture and discarding in summer temperatures has been shown to result in reduced survival (Chapman, 1981; Castro et al., in press). Further experiments, using higher temperatures, may be required to confirm the effects of elevated temperature.
Osmotic concentrations of the haemolymph of trawled Nephrops measured in the field were significantly lower than would be expected if animals had been sampled directly on the sea bed where the seawater was 33.5 psu. It is clear that haemodilution (by electrolyte depletion) had taken place during the haul. Surprisingly, no further lowering of blood osmotic concentration had occurred by the end of the 90 min emersion period on deck. In fact, some increase in osmotic concentration and K+ was observed, probably due to dehydration. No attempt was made to assess the condition of the animals at this stage. However, it is clear that the changes in body fluid concentration and composition are rapid and can occur during the upward passage of Nephrops through the surface layers (average salinity = 17.5), within the cod-end and during sorting on deck. Further work needs to be done to identify critical stages in the haemodilution process.
Control animals (CSW and CEM) tested for endurance in terms of the total number of tail-flips obtained by exhaustive stimulation showed strong responses at the end of the experiments. Although not tested at intermediate time intervals, it is clear by observation that, in the period immediately following low salinity exposure (from 48 h onward), Nephrops exposed to low salinities will have tail-flip performances which would be significantly reduced. In addition, animals subjected to low salinity exposure showed reduced capacity at the end of the experiment, even though they appeared quite normal in other respects. Surprisingly, animals exposed continuously to 15 seawater were less affected. Following discarding through this low salinity, some individuals developed a characteristic whitish "opaque" abdomen quite different from its normal translucent appearance. At the end of the experiment these animals developed melanization anterior to the uropods, and an absence of the normal tail-flipping response when tested. Similar symptoms have been observed in individuals following capture off the west coast of Scotland (Wileman et al., 1999; Stentiford and Neil, 2000). The latter authors have shown that this condition is due to a necrosis of the abdominal musculature and have suggested that periods of rapid, repetitive tail-flipping occurring during capture and handling may be the cause of these symptoms. It is possible that some of the individuals in our experiments showing reduced tail-flipping performance may be suffering from limited muscle damage while not showing major areas of necrosis of the deep abdominal flexor muscle. However, the possibility that swelling of muscle tissues due to osmotic uptake of water caused by rapid dilution of the extracellular fluid may also contribute to this cell damage should not be discounted since activity levels in our animals were deliberately maintained at relatively low levels during the experiments yet similar symptoms developed in many individuals.
In conclusion, simulation of capture and discarding through low salinity surface layers shows that exposure to this additional stressor results in increased mortality, significant disturbance of escape behaviour, and changes to physiological functioning which may affect tail-flip endurance and frequency compared with animals discarded through full-strength seawater in the upper layers. The extent of mortality observed here will be additional to those reported for Nephrops fisheries where low salinity stress was not apparent.
If it is assumed that the discard mortality of trawl-caught Nephrops under commercial conditions is 75%, and that it is 0% for creel-caught animals (Wileman et al., 1999), the stock management implications of the additional mortality can be calculated. The trawl mesh size currently being used in the Skagerrak–Kattegat fishery (70 mm) and the relatively high minimum landing size of 40 mm carapace length results in a large proportion of trawl-caught Nephrops being discarded as undersized back into the sea. The impact of an average additional mortality of 31% (range 23–39%) caused by a lower salinity surface layer (15 psu) was estimated by applying this mortality factor to the survivors of trawling (25%), and on all discarded sizes of creel-caught Nephrops since they are assumed to survive the fishing treatment if not exposed to low salinity surface water. Escape survival was assumed to be 90% for trawl-caught and 100% for creel-caught Nephrops (Wileman et al., 1999). The number of removals (NR) is calculated as
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Given the exploitation pattern of the Skagerrak–Kattegat trawl-caught Nephrops (ICES, 2001), and a surface salinity of about 15, the number and weight of removals from the Skagerrak–Kattegat stock is estimated to increase by, respectively, 6.3% (4.7–7.8%) and 4.0% (3.0–5.0%) (the values in brackets are 95% confidence limits). The corresponding figures for creel-caught Nephrops are 7.6% (5.7–9.5%) and 3.8% (2.9–4.8%), respectively.
Considering that the lowest surface salinities (15–18) occur in the Kattegat from April to July, this additional mortality will only have this level of impact during those months and will be reduced at other times of the year. In the Skagerrak, surface salinities do not fall so severely and the additional mortality is probably of minor importance in stock assessment there. Nevertheless, it suggests that the survival of undersized Nephrops in the fishery overall could be improved by making the fishing gear more size-selective, rather than requiring fishermen to size-sort the catch and discard the undersized back into sea thus exposing them to the effects of low salinity.
Apart from changing to more size-selective trawls, simple measures which could be taken to reduce the impact of low salinity water might include (a) reducing the time between encountering the low salinity layer (10–15 m below the surface) by faster hauling at the end of the tow; thus animals would have less time to ventilate their gill chambers with low salinity water, (b) reducing the time animals are held on deck, and (c) placing the discards in high salinity water during sorting and releasing them in this medium. It would be interesting to investigate the condition of discards subjected to such remedial treatments.
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R. R. Harris was in receipt of an EC Large Scale Facility Grant while working at Kristineberg Marine Research Station. We would like to thank the Director and staff for their assistance in carrying out this study. We also wish to thank the Director of the Fisheries Laboratory, Institute of Marine Research, Lysekil for his hospitality.
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