Not to be cited without prior reference to the authors
International Council for the
Exploration of the Sea
CM 2002/V:04
Discarding Norway lobster (Nephrops norvegicus) through low
salinity layers - mortality and damage seen in simulation
experiments.
R. R. Harris * and M. Ulmestrand **
* Department of Biology, University of Leicester, Leicester LE17RH, U.K. [tel +44 (0) 116
2523341: fax +44 (0) 116 2523330: e-mail: [email protected]].
** Institute of Marine Research, P.O.Box 4, 453 21 Lysekil, Sweden [tel. +46 (0)523 187 00:
fax +46 (0)523 139 77: e-mail: [email protected]].
Abstract
The Kattegat/Skagerrack Nephrops fishery is unusual in that animals normally live in high
salinities (33-34‰) but are raised through a low salinity surface layer and, if discarded, descend
back through it to the seabed. 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. A lower limit of 29 - 30‰ for its distribution has been suggested. Using CTD
data from Anholt E and Fladen hydrographic stations, near to known Nephrops grounds, a
knowledge of hauling times (including washing the cod-end with surface water) and sinking
rates, 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 over a 4/5
day period. "Discarded" Nephrops experienced significant haemodilution and gained water
rapidly. Animals showed slow rates of "tail-flipping", or absence of responses to stimulation, in
the period immediately following return to 33‰ seawater, although many showed recovery
later. Delayed effects included abdominal stiffness, swelling and further mortalities (25 - 42%
overall). Controls (exposed to 33‰ seawater only) showed good survival and vigorous
responses, even with a period of emersion. The effects of salinity exposure, additional to the
stresses of being trawled, on the fitness of discarded animals is evaluated.
Introduction
The commercially valuable Norway lobster Nephrops norvegicus 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 (Chapman and Howard, 1988;
Farmer, 1975), 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 compared to the MLS of 25 mm in the North
Sea area. This implies that more than half of the average Nephrops catch in the
Skagerrak/Kattegat area is below MLS and isdiscarded back to 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, 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 (seabed 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
seabed.
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. Longperiods 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 fisheries are 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
1
and, if discarded, are allowed to descend back through it to the seabed. 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 not abundant. 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
in salinities very different from full-strength seawater (Mantel and Farmer, 1983; Schoffeniels
and Dandrifosse, 1994).
The aims of the present study was to assess the effects of low salinity exposure on discard
mortality and 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.
2
Materials and Methods
Small adult Nephrops generally falling within the size range of animals which would be
discarded in Nephrops trawl or creel fisheries (< 40 mm CL) (mean mass = 35.94 ± 0.90 g; n =
72), were caught by baited creels in the Skagerrak near the mouth of Gulmarsfjorden, West
Sweden (N 58o 15' E 11o 26') at about 40 - 55 m depth in April 1999. Field conditions were:
bottom salinity = 33.13 ± 0.09 ‰; bottom temperature = 5.14 ± 0.05 o C (mean ± SE; n= 15)
(data courtesy of Kristineberg Marine Biological Station). Animals were transferred to flowing
seawater (33 ‰; 5oC) in holding tanks at Kristineberg Biological Research Station,
Fiskebäckskil, Sweden. 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.
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 results 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 pieces of Mytilus flesh placed within each tube on alternate days. Each
experiment commenced with a period of about 46 h in 33‰ seawater when animals were unfed.
Groups of six Nephrops within tubes were placed in glass aquaria (60 x 30 x 30 cm) connected
to the cooled, circulating seawater supply in a temperature - controlled room (salinity = 33‰,
temperature = 5oC; contained volume of seawater = 27 L). Animals were removed at intervals
for weighing and haemolymph sampling. At this time the condition of each animal was also
visually assessed according to a scale of four conditions (see below).
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 created. We also included in the protocol a
3
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 seabed.
Choice of low salinity exposure times.
A knowledge of CTD data at the Anholt E (N 56°40’, E 12°07’) and Fladen (N 57°11’,
E 11°40’) sampling stations within the Kattegat (courtesy of Swedish Meteorological and
Hydrological Institute) were used to design the simulation experiments. These stations are near
to known Nephrops grounds. Figure 1a and b shows the mean monthly salinity during 1998 at
the surface and at 40 m depth at the Anholt E 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 the depth, between 25 and 80 fathoms (45 to 145 m), also varied). From
a knowledge of the extent of the low salinity layer, we calculated that trawled Nephrops would
spend a maximum of 6 minutes within it when being brought to the surface. This would include
a period of washing the cod-end, at the ship side, with surface water to remove sediment.
Subsequently animals would spend an average of 90 minutes on deck during sorting. 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‰ seawater, and at two temperatures (5 and 15oC). 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 5oC = 8.88 ± 0.27; at 15oC = 8.59 ± 0.22 m min-1). A
period of emersion produced a minor, but statistically non-significant, increase in sinking rate
(9.67 ± 0.82 and 8.82± 0.21). On the bases 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.
4
Groups of 12 animals were exposed to the following:
(a) Controls were held in the conditions referred to above in 33 ‰ seawater at 5.0 (± 1.0)o C for
120 h. Body masses were determined and haemolymph samples taken at 24, 48, 72, 96 h.
(b) Treatment 1 animals were held in 33 ‰ seawater at 5.0 ± 1.0o C for 46.67 h. The medium
was then replaced with 27 L of 15 ‰ seawater (diluted by addition of distilled water to full
strength seawater; 33‰) at 5.0o 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 ‰ seawater for 2 min, before being returned to 33 ‰ seawater for the
remainder of the experiment. Animals were sampled at the intervals described in (a)
(c) Treatment 2 animals were held in 33 ‰ seawater at 5.0o C for 46.67 h. The medium was
replaced with 15 ‰ seawater. Animals were held at 5.0o C for 98 min (1.63 h) continuously in
this medium and then returned to 33‰ seawater. There was no period of emersion. Animals
were sampled also at the same intervals as in (a).
(d) Treatment 3 animals were held in 33 ‰ seawater at 5.0o C for 46.67 h. The medium was
then replaced with 15 ‰ seawater at 15 (± 1)o C for 6 min. Animals were then placed. in their
tubes, into a plastic fish basket and held emersed in air at 15oC for 90 min. Following this they
were returned to 15 ‰ seawater at the same temperature for 2 min, and then replaced in 33 ‰
seawater at 5.0o C for the remainder of the experiment.
(e) Treatment 4 animals were treated similarly except that submersion in 15‰ was for 98 min
continuously, with no period of emersion in air; the whole period in 15‰ was at a temperature
of 15oC.
(f) A control - in - air group was also included in which animals were held in 33 ‰ seawater at
5.0o C for 46.67 h then placed in a fish basket for 90 min at 15o C, then returned to 33 ‰
seawater at 5.0o C.
For all treatments, visual assessments of condition were made at these times and, additionally,
at 120 h.
Animals were wet - weighed (± 0.1 g) after superficial water was removed and drained from the
gill chambers by gentle shaking the animal held in a head - down position. Successive
weighings of control animals showed that this procedure gave consistent results, with an error
5
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 pereiopod and transferred immediately to Eppendorf tubes and shaken vigorously
for 2 min. Sampling was generally completed in < 30 sec to minimise emersion and the stress of
handling. Samples were then centrifuged (x 5000g) to separate off clotting proteins and the
supernatant used for all analyses. Samples were stored frozen (-20o C) until required.
At the end of the experiments, animals were tested for tail - flip escape swimming performance
by methods adapted from Newland et al. (1988) and Field (1992). Animals were transferred
individually into an opaque plastic aquarium (0.9 x 0.5 x 0.5 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 tailflips (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.
During all experiments the condition of animals was examined at each sampling interval and the
following scale was used to indicate their physical state: Condition 1 showed active, strong tail flipping, Condition 2 showed slow tail-flipping, but with some appendage movement, Condition
3 showed complete abdominal immobility but with some reflexes in limbs when touched, and
Condition 4
animals showed no reflexes, nor any scaphognathite activity, and appeared
moribund or dead.
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 N 56o 47'68 : E 11o 51' 11 in the Kattegat on 15 September 1999. CTD profiles at
this station showed bottom and surface salinities averaging 33.5 and 17.5 ‰ respectively, and
temperatures of 11 and 17.5 o C, with a surface low salinity layer extending down to ~ 15 m
depth. Samples were removed by 1 ml disposable hypodermic syringe and frozen (- 15oC).
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 (20µl samples in both cases), and
6
osmotic concentration by freezing-point depression (Roebling micro-osmometer). [K+] was
measured by Atomic Absorption spectrophotometry (Varian AA6), following dilution (50µl in
10 ml deionised water) of standards (5-50 mmol/l) and haemolymph.
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.
Results
Animal mortality and condition following low salinity exposure
Control animals maintained throughout the test period in 33‰ seawater showed good survival,
even following a 90 min period of emersion (air exposure) at 15oC (control or Control - in - air
animals). However, in all treatment groups experiencing a 98 min period of exposure to 15‰,
either including emersion following a shorter period of low salinity exposure at 5oC (Treatment
1), or continuous submergence (Treatment 2) at this temperature, or following similar
treatments at 15oC (Treatments 3 and 4), 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. Using the above scale (Condition 1-4 ) to indicate
these changes, all treatments showed significant differences compared to controls (P < 0.001).
Figure 2 shows the number of animals in each group demonstrating each of these conditions at
each sampling interval.
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 to controls which consistently showed
vigorous, rapid rates of tail-flipping (Condition 1 ).
At 48 h there were no significant
differences in condition between Treatments 1- 4 (median conditions were 1.5, 2.0, 3.0 and 2.5,
respectively) with 58 to 83% of individuals showing Condition 3. At 72 h ( 24 h after return to
33‰ seawater), Treatment 1 and 2 animals (both at 5oC) showed some recovery with a rise in
the proportion of animals showing Condition 1 and 2 but some mortality occurred also. In
7
Treatments 3 and 4 a similar recovery was seen, but 17 - 25% of the group failed to survive and
were removed. Mortalities continued during the course of the experiment, finally reaching 42%
and 25% in Treatments 1 and 2 respectively, and 25 and 33% in 3 and 4.. In Treatments 3 and 4
a significantly larger proportion of individuals were in Condition 2 - 3 and fewer in Condition 1
at 120 h than was the case for other Treatments. A number of individuals developed a whitish,
opaque abdomen, in addition to showing an absence of tail-flip responses. Controls showed
good survival and the great majority showed vigorous tail-flipping (Condition 1) throughout the
experimental period.
Changes in body mass following low salinity exposure
Control animals held in 33‰ 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 have no important effects on
body mass under these conditions (Figure 3a). In contrast, animals experiencing low salinity
conditions (15‰), either with a period of emersion or by continuous submersion (Treatments 1
and 2, respectively), showed highly significant gains in mass (expressed as % original). In
Treatment 1 the mean mass increased to 102.5 ± 0.49 % of the original wet weight when
measured immediately after removal (48 h) (P < 0.001), while in Treatment 2 a similar
significant gain was seen (P < 0.01). Similarly Treatments 3 and 4 (low salinity at 15oC)
showed significant gains in mean mass following low salinity exposure. When comparing all
treatments, it was found the extents of the mass gain immediately following exposure were not
significantly different. Control - in - air animals showed a non - significant reduction in mean
body mass after emersion (Figure 3a & b).
After low salinity exposure all animals were returned to 33‰ seawater for a period of 48 h.
During this period animals in the majority of treatments showed restoration to initial levels. The
exceptions were: Treatment 1 which showed a significantly lower mean mass than initial at 72 h
(97.8 ± 0.47 % initial), and Treatment 4 where mass remained at 98.4 ± 0.39 % initial at 72 h.
At the termination of the experiments, surviving animals of all treatment showed no significant
differences in their masses compared to initial levels.
Changes in haemolymph solutes following low salinity exposure
8
Control animals held in 33‰ seawater throughout the experimental period maintained
haemolymph osmotic concentrations slightly, but significantly, hyposmotic (lower than) to
seawater (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‰ ( 457.9 ± 3.18 mOsm Kg-1) of Treatment 1 and 2
animals was followed by a rapid significant reduction in haemolymph osmotic concentration (P
< 0.001) (Figure 4a). Animals were not sampled immediately prior to the low salinity exposure
period to minimise 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. Treatment 1 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 control animals sampled at the same intervals (P < 0.001).
The mean haemolymph osmotic concentration recorded immediately post-treatment was not
significantly different from Treatment 2 animals (P > 0.05) in which continuous exposure to
15‰ was maintained throughout 98 min. Following a return to 33‰, haemolymph osmotic
concentrations in both Treatments 1 and 2 increased to levels not significantly different to those
of controls within 24 h. These levels were also maintained for the remainder of the experiment.
Similarly, Treatment 3 and 4 animals showed highly significant decreases in haemolymph
osmotic concentration following low salinity exposure (at 15o C, in contrast to Treatments 1 and
2 which were at 5oC) with average decreases of 138 and 164 mOsm Kg-1, respectively, being
observed (P < 0.001) (Figure 4b). There was no significant difference in haemolymph osmotic
concentrations in the two groups during recovery in 33‰. Thus, at both temperatures, being
emersed or submerged during the low salinity exposure period had no significant effect on the
degree of the haemodilution.
Following return to 33‰ seawater haemolymph osmotic
concentrations values in Treatment 3 and 4. animals also recovered to near original levels.
However in Control in air animals 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‰ seawater throughout the experimental period maintained a mean
haemolymph [Na+] concentration of 451.0 ± 4.5 m mol l-1 (n = 24), slightly hyperionic to
seawater (33‰ = 429.0 ± 4.9; 15‰ = 220.4 ± 7.4 m mol l-1 [Na+]; n = 8). Transfer to 15‰
seawater of Treatment 1 and 2 animals was followed by significantly large decreases in
9
haemolymph [Na+] (Figures 4c and d)(P < 0.001). Again, surprisingly, there was no significant
difference in haemolymph [Na+] between animals subjected to continuous exposure to 15‰
seawater and those briefly submerged, with an intervening period of emersion. The largest
decrease in [Na+] was seen in Treatment 3 where the mean [Na+] was 295 .07 ± 3.17 m mol l-1
at the end of the low salinity exposure period. Control in air animals showed a slight but
significant rise in haemolymph [Na+] (P < 0.01). On return to 33‰ all experimental groups
showed increases in haemolymph [Na+] to original levels, except Treatment 3 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 (Fig 4e and f). However there
were differences, compared to the total solute and [Na+] changes, in that the largest decrease in
[Cl-] was seen in Treatment 2 animals which had significantly reduced haemolymph [Cl-]
compared to Treatment 1 animals (emersed for the majority of the period of their low salinity
exposure). Treatment 3 and 4 animals showed lesser falls in haemolymph [Cl-]. In Control-inair animals haemolymph [Cl-] rose significantly above the initial levels and was higher than
Control animals (P < 0.01).
Haemolymph [K+] was maintained a level below that of the surrounding 33‰ seawater in
control animals (mean = 6.5 ± 0.2 m mol l-1; Medium = 11.5 ± 1.1 m mol l-1 [K+]; n = 8).
Transfer of Treatment 1 and 2 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 than controls in Treatment 1
and 2 animals ( P < 0.01) but these were not significantly different from each other. A similar
picture emerged in Treatment 3 and 4 (at 15o C) animals which showed even greater increases
in haemolymph [K+] (P < 0.01). In the case of Treatment 4 animals, a sustained elevation of
haemolymph [K+] was observed at 96 h which was about 3.5 fold higher than initial levels
(Figure 4g and h). It appears that following low salinity exposure at a higher temperature
haemolymph [K+] rose higher than levels seen in similar salinity conditions at 5oC.
The effect of low salinity on tail-flip swimming performance
Control animals tested with a minimum of disturbance at the end of the experimental period
showed an ability to perform a mean of 126.7 ± 5.1 (11) tail-flips to exhaustion (as defined
10
above). Following transfer to low salinity conditions there was a significant reduction in
performance in surviving Treatment 1 animals (P < 0.001)(Table 1). However the mean number
of tail - flips seen in Treatment 2 animals (108.0 ± 28.76, n = 9) was not significantly lower
than that seen in controls. In some cases, surviving animals showed no tail - flips at all. These
were predominantly females. A similar picture emerged when Treatments 3 and 4 were tested
and compared with controls -in-air animals. Control - in- air animals showed strong tail flipping abilities (mean 133.4 ± 5.9 (12)) in spite of a 90 min period of emersion. However
significant reductions in Treatment 3 and 4 animals were seen (P < 0.05). No significant
differences in performance were seen when comparing Treatments 1 and 2, nor when 3 and 4
were similarly tested. The duration over which tail - flipping showed no major change
following low salinity exposure in Treatments 1,2 and 3. Thus the number of tail - flips was
reduced but these could be elicited at a lower frequency over a period of time similar to that of
control and control - in - air animals. The exception was the Treatment 4 group in which a
significant reduction in duration compared to controls was seen (P < 0.05).
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 mins before sampling ( air temperatures = 16.4 o C), are compared in
Table 2 with those of the control animals (in ~ 33 ‰ seawater) discussed above. The mean of
immediate post - capture haemolymph osmotic concentrations was 807.3 ± 11.6 mOsm Kg-1
(20). This was significantly lower than the levels shown by control animals, suggesting some
loss of solute from trawled animals (P < 0.001). The ambient deepwater salinity was recorded
simultaneously as ~ 33.5‰. Similarly, low haemolymph [Cl-] were seen (P < 0.001). However
haemolymph [Na+] was not significantly reduced and [K+] was significantly higher (P <0.001).
Comparisons were also made of immediate post-capture solute concentrations with those
present after 90 min emersion on deck. Although all showed some increase, only mean osmotic
concentration and [K+] were significantly so (P < 0.05).
11
Discussion
No attempt was made in this study to simulate the full stresses of capture 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, as used here, resulted in significant mortalities
immediately on removal from the discard simulation conditions and returned 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 major disturbances in
behaviour immediately post - exposure such as reduced tail-flipping on stimulation, slow
movements and swelling of the abdomen. Similar symptoms of low salinity exposure have
been reported in other stenohaline marine animals.
Dall (1974) reported that the lobster
Panulirus longipes abruptly transferred to 25‰ showed marked swelling, while populations of
the shrimp Crangon franciscorum exposed short - term to low salinities responded to stimuli
weakly (Shaner et al., 1985).
In some studies, mortality was described as being due to
"bursting". In some cases survivors restored their body volumes. Body volume increases
(generally measured as increases in mass) occur widely in stenohaline species. Maia squinado
showed an increase of about 4% in 4 h (Schwabe, 1933), while Libinia emarginata and Pugettia
producta showed increases of 0.46 and 1.4% in the 1st hour after transfer to 28‰ seawater
(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 that this 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
12
predation by fish and avian predators (Evans et al., 1994). This will have further impact on
discard survival.
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 (Kirschner, 1991) as shown in early studies
with the spider crab Maia squinado (Schwabe, 1933). 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 so that it was necessary to carry out some preliminary studies to obtain
basic data. Nephrops attained osmotic equilibrium in 25‰ (with 50% survival) and 28‰
seawater (100% survival), but in 21‰ seawater 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 than the
levels recorded from animals in 25‰ in the preliminary experiments by about 100 mOsm Kg-1
(difference in [Na+] = 52 ; [Cl-] = 63 m moles l-1). This suggests that during the brief low
salinity exposure simulation period, haemolymph concentrations had not fallen to some critical
low level since those animals which were maintained long - term in 25‰ showed lower values
(mean = 768 mOsm Kg-1 ). The rate of solute concentration decrease may be a factor
determining survival following a brief period of 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 Maia squinado subjected to hypoosmotic seawater. This effect, caused
13
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 the hyperpolarisation which was responsible for
reducing action potentials. 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 15oC. 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 are due to an
osmotic disturbance of excitable cell functioning remain to be determined.
Significantly, water content gain (as measured by mass) and haemolymph solute losses
occurred in Nephrops exposed to low salinities at rates which were similar 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 control - in - air
animals, emersed in air but maintained in 33‰ throughout, 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 fullstrength 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.
14
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 15oC, apart from
the steep rise in haemolymph [K+] seen in the latter group. However at the termination of the
experiments (120 h) Treatment 3 and 4 showed weaker states compared to Treatment 1 and 2
suggesting a damaging effect of the warmer temperature used. Nephrops is generally regarded
as a cold water species and capture and discarding in summer temperatures (25 - 30oC) has been
shown to result in reduced survival in the southern Portugal fishery (Castro et al., 2002).
Further experiments, using higher temperatures, may be required to confirm the effect of
elevated temperature.
Control animals showed strong responses at the end of the experiment when animals were
tested for their abilities in terms of total number of tail-flips obtained by exhaustive stimulation
(a measure of endurance). Although not tested at intermediate time intervals, it is clear by
observation that, in the period immediately following low salinity exposure (at 48 h), tail-flip
performance is significantly reduced. In addition, animals subjected to the simulation of
discarding involving 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 showed a less diminished performance. Following the
discard simulation in low salinities, some individuals developed a characteristic whitish
'opaque' abdomen quite different from the normal translucent appearance. At the end of the
experiment these animals developed melanisation 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 the necrosis of the abdominal
musculature and have suggested that period 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
15
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 to animals discarded through full-strength seawater upper
layers. The level of mortality observed here, which would be additional to those reported for
Nephrops fisheries where low salinity stress is not apparent (e.g 69%; Wileman et al., 1999),
would appear to indicate a serious problem for the management of the Kattegat/Skaggerak
fishery, particularly in Spring and early Summer. This estimated additional source of mortality
implies that nearly all discarded Nephrops will die in the Skagerrak-Kattegat Nephrops fishery.
Together with the significantly lower trawl net escape mortality estimated in Wileman et al.,
(1999), this underlines the need for a change to more size selective trawls in this fishery. It
would have a positive effect on the survival of undersized Nephrops, and it would likely
increase long-term yield in landings (ICES, 1997).
Osmotic concentrations of the haemolymph of trawled Nephrops measured in the field showed
significantly lower concentrations than would be expected if animals had been sampled directly
on the seabed where the seawater was ~ 33.5‰. 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 in the upward
passage 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. 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 reduced time to ventilate their gill chambers with low salinity
water (b) reducing the time animals are held on deck (c)place the discards in high salinity water
during sorting and discard them in this medium. It would be interesting to investigate the
16
condition of discards subjected to remedial treatments to see if the symptoms of low salinity
exposure are ameliorated.
Acknowledgements
Dr. 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,
17
Lysekil
for
his
hospitality.
References
Bergmann. M. J. N., Ball, B., Bijleveld, C., Craeymeersch, J. A., Munday, B. W., Rumohr, H.,
and van Santbrink, J. W. (1998) Direct mortality due to trawling. NIOZ-Rapport 1998-1
RIVO-DLO Report C003/98, Netherlands Institute for Sea Research, Den Burg, Texel,
404p.
Bergmann, M. and Moore P.G. (2001) Survival of decapod crustaceans discarded in the
Nephrops fishery of the Clyde Sea area, Scotland. ICES Journal of Marine Science, 58:
163 - 171.
Castro, M., Araújo, A., Monteiro, P., Madeira, A. M. and Silvert, W.(2002) The efficacy of
releasing caught Nephrops as a management measure. Fisheries Research (In Press).
Chapman, C. J. Shelton, P. M. J., Shanks, A. M. and Gaten, E. (2000). Survival and growth of
the Norway lobster Nephrops norvegicus in relation to light-induced eye damage.
Marine Biology, 136: 233-241.
Chapman, C. J. and Howard, F. G. (1988) Environmental influences on Norway lobster
(Nephrops norvegicus) populations and their implications for fishery management.
Symposium of the Zoological Society of London, 59: 343-353
Cornell, J. C. (1980) Salt and water balance in two marine spider crabs, Libinia emarginata and
Pugettia producta. III Some factors involved in short-term adaptations to a dilute
medium. Biological Bulletin (Woods Hole), 158: 16-25.
Dall, W. (1974) Osmotic and ionic regulation in the Western Rock Lobster Panulirus longipes
(Milne-Edwards). Journal of Experimental Marine Biology and Ecology, 15: 97-125.
Davenport, J. (1985) Osmoregulation in Marine Animals. In Physiology of Marine Animals Ed
by M. Laverack, Society for Experimental Biology Symposium No. 39. Cambridge
University Press, Cambridge.
Evans, S. M., Hunter, J. E., Elizal, R. I. And Wahju, R. I. (1994) Composition and fate of the
catch and bycatch in the Farne Deep (North Sea) Nephrops fishery. ICES Journal of
Marine Science, 51: 155-168.
Farmer, A. S. D. (1975) Synopsis of Biological data on the Norway Lobster Nephrops
norvegicus (Linnaeus, 1758). FAO Fisheries Synopsis No. 12, FAO Rome,97p.
Field, R. H. (1992) The control of escape behaviour in, and the physiology of, the Norway
Lobster, Nephrops norvegicus (L.). PhD Thesis, University of Glasgow.
18
ICES, (1997) Report of the Working Group on Nephrops Stocks. ICES CM 1997/Assess:11.
Kirschner, L. B. (1991) Water and Ions. In Environmental and Metabolic Animal Physiology 4th
Edition. Ed. by C. Ladd Prosser, Wiley-Liss, New York, Chichester, Brisbane, Toronto,
Singapore, 84p.
Lignon, J. M. and Gendner, J-P., (1988) Perméabilité cuticulaire branchial aux ions chez les
Crustacés décapodes et salinité du milieu environnant. Océanis, 14: 487-503.
Mantel, L. H. and Farmer, L. L. (1983) Osmotic and Ionic Regulation. In Biology of the
Crustacea Vol. 5. Ed. by D. E. Bliss, Academic Press, p54-161.
Newland, P. L., Chapman, C. J. and Neil, D. M. (1988) Swimming performance and endurance
of the Norway Lobster Nephrops norvegicus. Marine Biology 98: 345-350.
Poulsen, E. M. (1946) Investigations on the Danish fishery for, and the biology of, the Norway
lobster and the deep sea prawn. Report of the Danish Biological Station. 48: 27-49.
Redant, F and Polet, H (1994) Results of discard study on the Belgian Nephrops fishery in
central North sea. ICES C. M. 1994/K:44.
Robertson, J. D. (1949) Ionic regulation in some marine invertebrates. Journal of Experimental
Biology, 26: 182-200.
Robertson, J. D. (1960) Osmotic and Ionic Regulation. In The Physiology of Crustacea. Vol. 1
Ed. by T. H. Waterman, Academic Press, New York and London.
Robertson, J. D. (1961) Studies on the chemical composition of muscle tissue. 2. The
abdominal flexor muscles on the lobster Nephrops norvegicus (L.). Journal of
Experimental Biology 38: 707-728.
Schwabe, E. (1933) Über die osmoregulation verschiedener krebse (malacostracen). Zeitschrift
Vergleichender Physiologie, 19: 183-236.
Schoffeniels, E. and Dandrifosse, G. (1994) Osmorégulation: aspects morphologiques et
biochimiques. In Traité de Zoologie. Crustacés. Tome VII Fascicule 1. Ed. by P-P
Grassé, Masson, Paris.
Shaner, S. W., Crowe, J. H. and Knight, A. W. (1985) Long - term adaptation to low salinities
in the euryhaline shrimp Crangon franciscorum (Stimpson). Journal of Experimental
Zoology, 235: 315 - 323.
Sokal, R. R. and Rohlf, F. J. (1995) Biometry - the principles and practice of statistics in
biological research. 3rd edition. W. H. Freeman, New York. 887 pp.
19
Spicer, J. I., Hill, A. D., Taylor, A. C. and Strang, R. H. C. (1990) Effect of aerial exposure on
concentration of selected metabolites in blood of Norwegian lobster Nephrops
norvegicus (crustacea: Nephropoidea). Marine Biology, 105: 129 - 135.
Stentiford, G. D. and Neil, D. M. (2000) A rapid onset, post-capture muscle necrosis in the
Norway lobster, Nephrops norvegicus, (L.), from the West coast of Scotland. Journal of
Fish Diseases, 23: 251-263.
Symonds, D. J. and Simpson, A. C. (1971) The survival of small Nephrops returned to the sea
during commercial fishing. Journal du Conseil International pour l'Exploration de la
Mer, 34: 89-98.
Treherne, J. E. (1980) Neuronal adaptations to osmotic stress. Comparative Biochemsitry and
Physiology, 67B: 455-463.
Wileman, D. A., Sangster, G. I., Breen, M., Ulmestrand, M., Soldal, A. V. and Harris, R. R.
(1999) Roundfish and Nephrops survival after escape from commercial fishing gear.
Final report to the EC (FAIR-CT95-0753), Brussels.
Zainal, K. A. Y., Taylor, A. C. and Atkinson, R. J. A. (1992) The effect of temperature and
hypoxia on the respiratory physiology of the squat lobster Munida rugosa and Munida
sarsi (Anomura, Galatheidae). Comparative Biochemistry and Physiology, 101A: 557567.
20
Table 1. Tail-flip swimming performance in Nephrops, measured as total number of tail-flips
obtained by exhaustive stimulation, and time period over which response could be elicited.
Under Group, 1 - 4 refer to Treatments. Means ± 1 SE (n = 6 - 12). a - e indicate means
significantly different from one another (P < 0.05).
Group
Total no.of tailflips
Duration of
response (sec)
Control
126.7 ± 5.1 (11)a
92.5 ± 3.8
Control - in - air
133.4 ± 5.9 (12)bc
114.0 ± 5.0de
1
86.3 ± 5.9 (7)a
117 ± 10.1
2
108 .0 ± 28.8 (8)
95.0 ± 6.1
3
84.9 ± 9.5 (9)b
89.7 ± 9.5d
4
69.0 ± 23.7 (7)c
65.9 ± 17.1e
21
Table 2. Haemolymph osmotic concentrations, [Na+], [Cl-] and [K+] of Nephrops trawled from
about 40 m depth at a station N 56o 47'68 : E 11o 51' 11 in the Kattegat on 15 September 1999.
Measurements of immediate post-capture and after 90 min emersion are show together (n = 20)
with those of Controls in 33‰ (n = 24). Deep and surface seawater analyses are also shown.
Means ± 1 SE. a - d indicate means significantly different from one another (P < 0.05).
Group
Osmotic
concentration
(mOsm Kg-1)
[Na+]
m mol l-1
[Cl-]
m mol l-1
[K+]
m mol l-1
Controls in
33‰
949.5 ± 2.0
(24) a
451.0 ± 9.6
437.5 ± 3.8 b
6.5 ± 0.2
33‰ seawater
958.1 ± 3.5
(8)
429.0 ± 4.9
457.2 ± 11.5
11.5 ± 1.1
Immediate
Postcapture
807.3 ± 11.6
(20) ac
427.7 ± 8.5
382.3 ± 8.1 b
11.3 ± 0.5 d
After 90 min
emersion
841.5 ± 17.2
(20) c
430.8 ± 11.0
393.0 ± 13.0
16.2 ± 2.1 d
Surface
seawater
504.4 ± 13.4
(3)
242.97 ± 8.0
252.4 ± 3.4
5.7 ± 1.2
22
Figure 1. (a) Map showing Kattegat and Skaggerak with Anholt E and Fladen sampling stations.
Typical seasonal changes in salinity at surface and 40 m depth (b) and a profile showing salinity
changes with depth (for April) at Anholt E (c)(courtesy of the SwedishMeteorological and
Hydrological Insitute).
a)
b)
23
c)
Figure 2. Condition of control Nephrops and in Treatments 1 to 4 following low salinity
exposure. Details of each Treatment is given in Materials and Methods. Legend indicates
shading used to indicate Condition 1-4 . (n = 12 in each Treatment).
control
No. animals in conditions 1-4
No. animals in condition 1-4
15 Treatment 1
No. animals in condition 1-4
Condition
15
15 Treatment 3
15
10
10
5
5
15
1
10
2
control in
air
10
3
5
5
4
0
48
72
96
0
120
48
15
10
72
96
120
96
120
96
120
Treatment 2
10
5
5
0
48
72
96
0
120
48
0
48
72
96
72
Treatment 4
0
120
48
Time of transfer (h)
72
Time of transfer (h)
24
Figure 3. Changes in body mass of Nephrops before and after low salinity exposure (Treatments
1 to 4). (a) Treatments 1 and 2 with Controls in 33‰ seawater (b) Treatments 3 and 4 with
Controls - in - air at 15oC (open square brackets). Treatments 1 - 4 are indicated on each graph.
* indicates significant difference between Treatment means and Controls , and ** significant
difference between Treatments. Means ± 1 SE (n = 12).
(a)
15 ppt
Body mass as % original
105.0
*
*
102.5
control
100.0
1
2
*
97.5
95.0
0
20
40
60
80
100
Time (h)
(b)
15ppt
emersed
Body mass as % original
105.0
*
*
102.5
3
4
100.0
*
control-inair
97.5
95.0
0
20
40
60
80
Time (h)
25
100
Figure 4. Changes in Nephrops haemolymph osmotic concentrations (a and b), [Na+] (c and d),
[Cl-] (e and f) and [K+] (g and h). Solid vertical lines indicate the period of each Treatment
which is followed by a recovery period in 33‰ seawater.Treatments 1 - 4 are indicated on each
graph. * indicates significant difference between Treatment means and Controls , and **
significant difference between Treatments. Means ± 1 SE (n = 12).
(a)
1000
Control
900
1 *
2 *
800
700
20
(c)
40
60
Time (h)
80
900
*
*
800
100
0
15 ppt
-1 )
500
450
control
400
2 *
1 *
350
300
250
0
20
40
60
20
80
500
(e)
4
control
-in-air
400
3
350
*
300
**
20
40
60
80
100
Time (h)
450
2
control
400
*
**
-1 )
1
15 ppt
emersed
(f)
500
*
3
Haemolymph [Cl](m moles l
-1 )
Haemolymph [Cl] (mmoles l
100
450
0
300
450
4
controlin-air
400
350
300
0
20
40
60
80
100
0
20
Time (h)
(g)
40
60
80
100
Time (h)
15 ppt
emersed
(h)
15 ppt
25
20
15
10
1
2
5
control
0
Haemolymph [K](m moles l
-1 )
-1 )
25
Haemolymph [K](m moles l
80
250
100
15 ppt
350
60
15 ppt
emersed
*
(d)
Time (h)
500
40
Time (h)
Haemolymph [Na] ( m moles l
-1 )
**
4
3
control
-in-air
700
0
Haemolymph [Na] ( m moles l
15 ppt
emersed
(b)
15 ppt
Haemolymph OC (mOsm Kg-1)
Haemolymph OC (mOsm Kg -1 )
1000
4
*
20
*
15
*
3
10
control
5
0
0
20
40
60
80
100
0
Time (h)
26
20
40
60
Time (h)
80
100