Alkaline tide and nitrogen conservation after feeding in an elasmobranch (Squalus acanthias)
1 Department of Biology, McMaster University, 1280 Main St West, Hamilton,
Ontario, Canada L8S 4K1
2 Rosenstiel School of Marine and Atmospheric Sciences, University of Miami,
Miami, Florida 33149, USA
3 Bamfield Marine Sciences Centre, 100 Pachena Drive, Bamfield, British
Columbia, Canada V0R 1B0
4 Department of Biology, University of Victoria, Victoria, British Columbia,
Canada V8W 2N5
* Author for correspondence (e-mail: woodcm{at}mcmaster.ca)
Accepted 10 May 2005
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Summary |
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Key words: gastric acid secretion, metabolic alkalosis, ammonia, urea, osmolality, shark
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Introduction |
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With this background in mind, we pursued the following objectives with the
Pacific spiny dogfish, Squalus acanthias. The first was to develop a
stomach tube feeding model because dogfish will not eat naturally when
confined in small boxes, as required for the chronic blood sampling of
acidbase studies. The second was to establish whether an alkaline tide
occurs after dogfish are fed a realistic ration by this method and, if it
does, whether it is modulated by `respiratory compensation'
(CO2 retention to limit pH
increases), as in many higher vertebrate carnivores (reviewed by
Andrade et al., 2004
). The
third was to determine the effects of this feeding regimen on nitrogen
metabolism, with particular emphasis on ammonia-N and urea-N excretion rates
and corresponding blood levels of these two end products. The final goal was
to determine changes in associated parameters (blood
O2, glucose, trimethylamine
oxide, total amino acids, creatinine, osmolality, chloride, glucose,
ß-hydroxybutyrate) diagnostic of possible osmoregulatory and metabolic
phenomena associated with feeding.
Our results demonstrate that feeding causes disturbances in many of the measured parameters, particularly a marked, uncompensated alkaline tide, and reinforce the view that dogfish are strongly N-limited, excreting little `excess' ammonia-N and no `excess' urea-N after a meal. The present study forms part of a two-phase investigation on this topic, the other component focussing on some of these same parameters in naturally fed, non-catheterized dogfish and on the accompanying enzymatic changes in various tissues (M.K., P.J.W., T.P.M. and C.M.W., unpublished results).
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Materials and methods |
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Each dogfish was anaesthetized with MS-222 (0.2 g l1), placed on an operating table, weighed, and fitted with indwelling catheters. In Series 1, the catheters consisted of a stomach feeding tube, a caudal artery cannula and a caudal vein cannula, and the anus was tied off to prevent defecation on the supposition that the latter might confound flux measurements (but see below). The anus was ligated with silk suture just anterior to the rectal gland, through a 3 cm incision in the ventral midline In Series 2, only a stomach feeding tube and a caudal artery cannula were inserted, and the anus was not ligated. This change in Series 2 (lack of anal ligation) was adopted because N-excretion rates, particularly those of urea-N, were high and very variable in Series 1.
After Series 1 was completed, a subsequent pilot series with uncannulated
animals revealed that anal ligation caused about a 50% increase in urea-N
excretion rate, as well as greater variability in both ammonia-N and urea-N
excretion rates, probably explained by stress and/or minor blood loss from the
incision site. We also noticed that struggling sometimes resulted in fin
abrasion against the walls of the fish chambers, another potential source of
minor blood loss. Indeed, given the very high levels of urea-N in the
bloodstream, blood loss of only 0.3 ml kg1
h1 would be required to cause this elevation. Therefore,
only the flux data of Series 2 have been reported, where anal ligation was not
employed, and particular care was taken to minimise disturbance.
Based on a number of pilot tests, the final stomach tube design used in
both series was individually fitted to each fish via the oesophagus,
terminating several centimetres anterior to the pylorus. This placement was
estimated based on the total length of the fish and was confirmed by autopsy.
The key point to successful placement was tailoring the length of the tube to
the size of the fish, which we did based on fitting tubes experimentally to a
number of dead animals of different lengths. The tube consisted of flexible
polyethylene tubing (0.32 cm internal diameter), which was heat polished at
the stomach end and heat moulded so as to allow its exit via a small
puncture wound through the jaw muscle at the side of the mouth and then firmly
ligated with silk suture at this point and along the upper jaw, terminating in
an upward projection of 3 cm anterior to the eye. Prior to insertion, the
tube was filled with 140 mmol l1 NaCl, the vehicle for the
food slurry (see below), and sealed with a plug at the anterior end. Caudal
artery (both Series 1 and 2) and caudal vein catheters (Series 1 only) made of
Clay-Adams (Becton-Dickinson, Sparks, MD, USA) polyethylene PE50 tubing were
implanted through a small hole in the haemal canal via a 5 cm
incision through the muscle of the caudal peduncle, as described by DeBoeck et
al. (2001
). The catheters were
filled with heparinised dogfish saline [sodium heparin, 50 i.u.
ml1; saline recipe as in Wood et al.
(1994
) but with urea level
raised to 400 mmol l1=800 mmol l1 urea-N].
Wounds were dusted with powdered oxytetracycline to avoid infection, tightly
closed with silk ligatures, and sealed with a sheet of rubber dental dam,
which was glued to the skin using tissue cement (3M Vetbond, 3M Animal Care
Products, St Paul, MN, USA).
After surgery, the dogfish were revived in anaesthetic-free water and
transferred to covered wooden fish boxes, which were coated with polyurethane,
the same boxes as used in an earlier study
(Wood et al., 1995). The boxes
were 105 cm in length, 16.5 cm in width and 25 cm in height, with a
flow-through of 1 l min1. Perimeter aeration over the
complete length of the box ensured good mixing during flux measurements. The
boxes were bathed in an external running seawater bath to maintain temperature
(1112°C) when flow-through was suspended for the flux measurements.
A recovery period of at least 36 h was allowed before experiments were
started.
Preparation and administration of food
Fillets of white muscle were cut from 12 freshly caught flatfish
(Hippoglossoides elassodon and Parophrys vetulus), two of
the common species in the trawl-caught fish that were routinely fed to the
dogfish in the main holding tank. The fillets were ground to a fine paste in a
Waring food blender, then stored frozen at 20°C in small samples
until used. A sample was taken for N-analysis. The `meal' administered to
experimental animals consisted of 2% of the dogfish's body mass of the
flatfish muscle paste mixed 50:50 with an equal volume of 140 mmol
l1 NaCl to create a smooth slurry that could be infused
via the stomach feeding tube. The rationale for using this vehicle is
that it would be representative of the teleost body fluids that are ingested
along with muscle when dogfish feed naturally. Thus, the volume infused
represented 4% of the body mass. From gut content measurements on naturally
feeding dogfish in the large tank, we established that this is well within the
range that dogfish can consume on a single feeding event (M.K., P.J.W., T.P.M.
and C.M.W., unpublished results). Control animals received 4% of the body mass
as 140 mmol l1 NaCl. For feeding, the meal was administered
as a bolus down the stomach tube over an approximate 5 min period. A small
extra volume of saline was used to flush the tube, which was then re-sealed
with a plug.
Flux measurements
For flux measurements, flow-through to the box was stopped and the water
level set to a mark representing 32 litres. Water samples (10 ml) for
ammonia-N and urea-N measurements were taken at the start and end of each flux
period. At times of water renewal (`flushes'), the water level was lowered to
the point where the animal's dorsal fin was just exposed, then filled to the
top of the box with fresh seawater, a procedure that was repeated three times
before the volume was reset to 32 litres.
Blood sampling
Blood samples were withdrawn via the catheters into ice-cold
gas-tight Hamilton syringes. In Series 1, arterial samples were taken prior to
venous samples, and two separate syringes of 0.6 ml and 0.4 ml were taken for
each. The first was used for blood gas and pH measurements, and the second was
immediately centrifuged (2 min at 9000 g), then the plasma was
divided into aliquot samples and frozen (80°C) for metabolite
measurements. In Series 1, blood recovered from the electrodes was mixed with
the red cell pellet, made up to the original 1.2 ml volume with
non-heparinised saline, and re-infused. In Series 2, there were no blood gas
measurements but a larger set of metabolite measurements was required, so a
volume of 0.9 ml was taken and immediately replaced with an equal volume of
non-heparinised saline; the red cells were not re-infused, so as to minimize
disturbance. Plasma was obtained by immediate centrifugation, then divided
into four tubes and frozen as above for various metabolite assays.
Experimental series
In Series 1, flow-through to the box was stopped for a 46 h period
for a pre-feeding measurement of ammonia-N and urea-N flux rates, after which
a set of pre-feeding blood samples was taken. A meal of either flatfish muscle
slurry (experimental animals, N=5) or saline (control animals,
N=5) was then administered via the stomach tube, the box
immediately flushed, and a new flux measurement started (0 h). Blood samples
were then taken at 1 h, 2 h, 3 h, 4 h, 6 h, 9 h, 1518 h (nominally 17
h), 24 h and 48 h post-feeding. Water samples were taken at every blood
sampling time, with flushes at 9 h, 17 h, 24 h, 36 h and 48 h. Blood samples
were analysed for blood gases and pH and plasma ammonia-N, urea-N, glucose and
chloride concentrations.
In Series 2, a similar protocol was used but with fewer blood and water samplings; particular care was taken to minimize disturbance. The same feeding methods as in Series 1 were employed for the experimental (N=8) and control (N=8) animals. Blood samples were taken after the pre-feeding flux period, prior to feeding, and at 2 h, 4 h, 6 h, 9 h, 18 h, 24 h, 34 h and 45 h post-feeding. Water samples were taken simultaneously, with flushes at 9 h, 18 h, 34 h and 45 h. Blood samples were analysed for plasma ammonia-N, urea-N, trimethylamine oxide-N (TMAO-N), creatinine-N, total free amino acids (FAA-N), glucose, ß-hydroxybutyrate, chloride and osmolality.
Analytical techniques
Arterial and venous blood oxygen tensions
(aO2,
vO2) and pHs (pHa, pHv)
were measured using Radiometer electrodes kept at the experimental temperature
with water jackets. True plasma CO2 was measured by the method of
Cameron (1971
) on plasma
obtained from blood samples centrifuged in sealed tubes. Outputs of the
electrodes (E5036 for
O2;
GK401C for pH; E5046 for
CO2 in the Cameron chamber)
were displayed on Radiometer pHM 71 and pHM 72 acidbase analysers.
Arterial and venous blood carbon dioxide tensions
(
aCO2,
vCO2) and bicarbonate
concentrations ([HCO3]a,
[HCO3]v) were calculated using the solubility of
carbon dioxide (
CO2), the apparent pK (pKapp) for
dogfish plasma, and rearrangements of the HendersonHasselbalch equation
according to Boutilier et al.
(1984
).
Plasma ammonia-N was measured enzymatically (L-glutamate
dehydrogenase; Raichem Ammonia Reagent; Product No. 85446;
Mondzac et al., 1965) on the
first thaw of frozen plasma. Plasma total free amino acid levels (FAA-N) were
measured using the ninhydrin assay (Moore,
1968
), with subtraction of previously measured ammonia-N, owing to
the partial detection of ammonia by the ninhydrin method
(Kajimura et al., 2004
). The
correction was small, because plasma ammonia-N concentrations were less than
3% of FAA-N concentrations. Plasma urea-N was measured with the diacetyl
monoxime method (Rahmatullah and Boyde,
1980
). Plasma TMAO-N levels were assayed by the ferrous sulphate
and EDTA method (Wekell and Barnett,
1991
). Plasma glucose was determined with hexokinase reagent
(Thermotrace kit 1542; ThermoElectron Corp., Waltham, MA, USA). Plasma
ß-hydroxybutyrate was measured enzymatically (ß-hydroxybutyrate
dehydrogenase) by the method of McMurray et al.
(1984
) using Stanbio
Laboratory kit 2440 (Boerne, TX, USA). Plasma creatinine-N was measured by the
colorimetric assay of Heinegarde and Tiderstrom
(1973
) (Sigma kit 555; Sigma,
St Louis, MO, USA). Plasma chloride was measured by coulometric titration
(Radiometer CMT-10; Copenhagen, NV, Denmark) and osmolality by vapour pressure
osmometry (Wescor 5100C; Westcor Inc., Logan, UT, USA).
Seawater ammonia-N and urea-N concentrations were determined by the
salicylate hypochlorite (Verdouw et al.,
1978) and diacetyl monoxime
(Rahmatullah and Boyde, 1980
)
methods, respectively.
The flatfish muscle paste (four replicates) was analysed for N-metabolites
in order to estimate the nitrogen load administered in a meal. The frozen
paste was pulverized in a mortar and pestle, which was chilled with liquid
nitrogen, then homogenized in 0.5 mol l1 HClO4
and centrifuged (4°C, 10 000 g, 5 min). Pellets were
washed twice with 0.5 mol l1 HClO4, then the
original and wash supernatants were pooled, neutralized with 3 mol
l1 KOH and then centrifuged (4°C, 10 000
g, 5 min) to remove precipitated KClO4. The pellet
was resuspended in 0.5 mol l1 KOH, incubated for 1 h at
37°C, then analysed for total protein content by the dye-binding assay of
Bradford (1976), using Sigma
reagent and bovine serum albumin as standard. A standard N-content of 0.16 g-N
g1 protein was assumed
(Cho, 1990
). The final pooled
supernatant from the washes was analysed for ammonia-N, urea-N, TMAO-N, FAA-N
and creatinine-N using the methods outlined above for plasma.
Statistics
Data are reported as means ± 1
S.E.M. (N). Pre-feeding measurements
were first compared using Student's two-tailed t-test (unpaired) to
ensure absence of pre-existing differences between experimental and control
groups (there were none). The post-feeding data were then subjected to two-way
(time, treatment) analysis of variance (ANOVA) to detect overall effects of
feeding. Student's t-tests (unpaired) were then applied to detect
specific differences between experimental and control groups at the same
sampling time, indicated by asterisks. One-way analysis of variance followed
by the LSD (least significant difference) test was applied to detect specific
differences within a treatment group (control or experimental); means not
sharing the same case letters are significantly different. Dunnett's paired
multiple comparison test was also employed to detect specific differences
within a treatment group (control or experimental), relative to the
pre-feeding value, and yielded the same results as those of the more
comprehensive LSD test, so only the latter are reported. A significance level
of 0.05 was used throughout.
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Results |
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Pre-feeding arterial
aCO2 (
1 torr) was
0.5 torr below
vCO2,
while
aO2 (
100 torr)
was much higher than
vO2
(
20 torr; note that 1 torr=0.1333 kPa). These parameters were again
unaffected by the loading of saline into the stomach (Figs
3A,
4A). Experimental feeding had
negligible influence on
aCO2 and
vCO2; the former was very
slightly elevated at 2 h only, and there were no overall effects of treatment
(two-way ANOVA), indicating a lack of respiratory compensation for the
alkaline tide (Fig. 3B).
aO2 also remained constant
at pre-feeding levels, but
vO2 declined by
50%
from 1 h to 6 h post-feeding (significant relative to the control group at 3 h
and 6 h; Fig. 4B), reflecting
greater O2 utilization from the blood.
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The influence of experimental feeding on ammonia-N and urea-N excretion
No differences were detected in N-excretion rates between the control and
experimental treatments in Series 1, but the rates, particularly those of
urea-N, were high and variable, probably due to minor blood loss associated
with anal ligation, struggling and/or fin abrasion. The protocol was modified
in Series 2 to avoid these problems (see Materials and methods). Figs
5 and
6 present data from those fish
of Series 2 (control N=5, experimental N=4) where there was
no struggling or potential for blood loss from fin abrasion. When data from
all the fish of Series 2 (control N=8, experimental N=8)
were analysed, trends and statistical significance were identical, although
variability was somewhat greater.
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Prior to feeding, urea-N excretion (750 µmol-N
kg1 h1) was approximately 13-fold greater
than ammonia-N excretion (
60 µmol-N kg1
h1). The administration of saline into the stomach of
control dogfish caused a small elevation in ammonia-N excretion at 02 h
(Fig. 5A). Experimental feeding
caused a slightly larger increase in ammonia-N excretion that was significant
at 02 h and also significantly greater overall relative to the control
treatment (two-way ANOVA), although there were no significant individual
differences between the two treatments at particular time points
(Fig. 5B). Over 45 h, the
elevation in ammonia-N excretion amounted to
700 µmol-N
kg1.
Urea-N excretion was unaffected by saline loading in the control group
apart from an unexplained increase in the latter at 69 h post-infusion
(Fig. 6A). Urea-N excretion was
not increased by experimental feeding. Overall urea-N excretion over 45 h was
actually lower in the experimental fish by 3200 µmol-N
kg1, although two-way ANOVA revealed no significant effect
of treatment (Fig. 6B).
The measured N-content in the meal of flatfish muscle
(Table 1) helps put these small
differences in ammonia-N and urea-N excretion into perspective. The total
N-content amounted to 1625 µmol-N g1 muscle, or 32
500 µmol-N kg1 dogfish, in the meal, more than 90% of
which was in the form of protein (Table
1). At the pre-feeding N excretion rates (ammonia-N=60 µmol-N
kg1 h1, urea-N=750 µmol-N
kg1 h1), the N-content of the meal would
be lost in
40 h.
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The influence of experimental feeding on plasma metabolites and ionoregulatory status
Absolute levels and responses of plasma metabolites were similar in Series
1 and 2, so the more extensive data set of the latter (arterial values) is
presented. Of the plasma parameters measured in Series 1 (ammonia-N, urea-N,
glucose, Cl), the only detectable arterialvenous
difference was in ammonia-N, which was approximately 50 µmol
l1 higher on the venous side.
Pre-feeding plasma ammonia-N levels were extremely low (150 µmol-N
l1) relative to plasma urea-N (
800 mmol-N
l1) and plasma TMAO-N levels (
70 mmol-N
l1; Fig. 7).
The administration of saline caused no change in plasma ammonia-N in the
control treatment but there was a persistent elevation in plasma ammonia-N
associated with experimental feeding, which was significant overall (two-way
ANOVA), with individual significant differences at 4 h, 18 h and 45 h
post-feeding (Fig. 7B). Plasma
urea-N levels tended to rise slightly in both treatments (significant only in
the control group at 18 h), but there was no overall effect of feeding
(two-way ANOVA) and no significant differences at any time between the two
treatments (Fig. 7A). Plasma
TMAO-N concentration was marginally affected by feeding, with a significantly
higher value at 4 h in the fed group but no overall effect (two-way ANOVA) or
other specific differences (Fig.
7C). Other plasma N-compounds (FAA-N
12 mmol-N
l1, creatinine-N
0.15 mmol-N l1), as
well as glucose (
6 mmol l1) and ß-hydroxybutyrate
(
4.5 mmol l1) were similarly little affected by the
treatments (Table 2). However
both FAA and ß-hydroxybutyrate were lower after feeding overall (by
two-way ANOVA), although there were no significant differences at specific
time points (Table 2).
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Plasma osmolality (940 mOsmol kg1) exhibited a clear
response to experimental feeding, with a post-feeding elevation that was
significant overall relative to the control treatment (two-way ANOVA), with
individual significant differences at 2 h, 4 h, 6 h, 9 h, 18 h and 34 h
(Fig. 8A). There was a slight
tendency for osmolality to decline in the control group. Post-feeding
differences averaged
25 mOsmol kg1 between the two
treatments. Changes in plasma Cl concentration (
255
mmol l1 prior to feeding) explained part of this difference
(Fig. 8B). Cl
tended to rise in the experimentally fed dogfish and fall initially in the
saline-loaded control animals. The difference was
10 mmol
l1, accounting for
40% of the osmolality difference,
and was highly significant overall (two-way ANOVA), although the only
individual significant difference was at 18 h.
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Discussion |
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Unlike members of other vertebrate classes (see
Andrade et al., 2004), the
dogfish exhibited virtually no evidence of respiratory compensation (i.e.
elevation of
CO2 to offset
the rise in pH; Fig. 3). The
likely reason for this difference is that the dogfish breathes exclusively
with its gills. Because of the much lower solubility of O2 relative
to CO2 in water, the gills of most water-breathing animals are
greatly hyperventilated with respect to CO2 excretion, such that
ventilatory adjustments have only a small effect on blood
CO2 levels
(Perry and Wood, 1989
).
Furthermore, although ventilation was not measured in the present study, it
was our impression that the dogfish were ventilating more deeply after a meal,
during the period when
vO2
was decreased (Fig. 4B). This
would tend to enhance rather than inhibit CO2 excretion. The
dogfish gill is particularly efficient at CO2 excretion because the
HCO3 dehydration reaction is catalysed in part by
extracellular carbonic anhydrase, which is both present on the endothelial
surfaces of gill blood vessels and circulates freely in the blood plasma
(Wood et al., 1994
; Gilmour et
al., 1997
,
2001
;
Henry et al., 1997
;
Wilson et al., 2000
). It may
be virtually impossible for the dogfish to `retain CO2'. In future
studies, use of a K+-stimulated, H+-ATPase inhibitor
such as omeprazole (see Andrade et al.,
2004
) may prove informative to separate changes in acidbase
status resulting from augmented gastric acid secretion from those associated
with altered ventilation and metabolic rate.
While ventilatory control of acidbase status appears to be minimal,
the ability of elasmobranchs to correct acidbase disturbances
via ion versus acidic/basic equivalent exchange mechanisms
at the gills is well established (e.g.
Heisler, 1988). Their ability
to resist the alkalinizing influence of basic equivalent loading is
particularly impressive (Wood et al.,
1995
; Tresguerres et al.,
2005
). Thus while the alkaline tide had been corrected by 17 h
post-feeding (Figs 1,
2), this does not necessarily
mean that the gastric H+ secretion responsible for the phenomenon
had subsided, but rather that branchial base excretion mechanisms had fully
caught up with the rate of metabolic base addition to the blood by the
acid-secreting cells of the gastric mucosa. Indeed, autopsies revealed that
digestion continued right up to 48 h (and probably longer) and that some food
paste still remained in the stomach by this time. Interestingly, food remnants
had not reached the anus by this time, so initial concerns (Series 1) about
defecation contaminating N-flux measurements proved unfounded. A slow time
course (relative to that of teleosts) for digestion, assimilation and gut
passage appears to be characteristic of elasmobranchs
(Jones and Geen, 1977
;
Wetherbee et al., 1987
;
Schurdak and Gruber, 1989
;
Wetherbee and Gruber, 1990
;
Cortes and Gruber, 1990
;
Sims et al., 1996
).
Using several approaches, it is possible to make a rough estimate of the
rate at which metabolic base was added to the blood by the acid-secreting
cells of the stomach. One approach is simply by comparison to the recent work
on S. acanthias of Tresguerres et al.
(2005), who found that a
HCO3 infusion rate of 1000 µmol
kg1 h1 induced a slightly larger metabolic
alkalosis than seen in the present study (Figs
1,
2). Another approach is to
employ the traditional technique pioneered by Rune
(1965
,
1966
) and now widely used in
humans (Niv and Fraser, 2002
),
whereby the rate at which `base excess' is generated in the blood is measured,
factored by a distribution space (0.3 of body mass). Applying this method, and
a blood non-HCO3 buffer capacity of 9 slykes (9
µmol pH unit1 g1) for S.
acanthias (Lenfant and Johansen,
1966
), to the data of Figs
1 and
2 from hours 14
post-feeding yields a rate of
475 µmol kg1
h1. This is undoubtedly an underestimate as some metabolic
base excretion across the gills would have already started during this period
(Wood et al., 1995
). Thus, the
rate of endogenous metabolic base generation associated with digesting the
experimental meal in the dogfish was in the range of 4751000 µmol
kg1 h1. By way of comparison, the reported
rate in human subjects after a normal meal of mixed composition is
425
µmol kg1 h1
(Rune, 1966
). It would be
interesting to test whether an even larger experimental meal would cause a
larger alkaline tide in the dogfish, or whether a maximal rate has been
achieved in the present study.
The literature is unclear with respect to the pattern of gastric acid
secretion in carnivorous elasmobranchs. Some reports
(Sullivan, 1905;
Caira and Jolitz, 1989
)
indicate that stomach pH is close to neutrality in fasting animals, with much
lower values in fed animals, including S. acanthias
(Sullivan, 1905
), in accord
with the present results indicating a sharp increase in secretion after
feeding. Others have reported a markedly acidic gastric pH regardless of the
presence or absence of food in the stomach
(Babkin et al., 1935
;
Menon and Kewalramani, 1959
;
Williams, 1970). A recent detailed study (Papastamiatou and Lowe, 2004) using
indwelling, continuously recording pH sensors in the stomachs of naturally
feeding leopard sharks (Triakis semifasciata) reported an extremely
acid stomach (pH
1.5) in starved animals, while feeding was associated
with a sharp rise in pH (to
3.5) followed by a progressive slow
acidification thereafter. The data were interpreted as reflecting a continuous
low level of acid secretion during fasting, which was accelerated after
feeding. The buffering action of the food was responsible for the initial
rapid rise in pH and for retarding the later fall in pH. It seems probable
that all elasmobranchs increase gastric acid secretion after feeding but that
there may be inter-species differences in whether it is completely turned off
between meals. In future studies, it will be of interest to track the pH and
buffer capacity of the stomach contents over time after feeding in S.
acanthias to evaluate the magnitude and temporal relationship between the
rate of gastric acid secretion and the alkaline tide in the blood.
Nitrogen conservation after feeding
Experimental feeding of the ureotelic dogfish resulted in no increase in
urea-N excretion (indeed, a non-significant decrease;
Fig. 6) and only a very small
rise in ammonia-N excretion (Fig.
5), amounting to less than 3% of the total-N in the meal. The
increase in ammonia-N excretion was accompanied by a modest increase in plasma
ammonia-N concentration (Fig.
7B). This is very different from the situation in ammoniotelic
teleosts, where ammonia-N excretion and plasma ammonia-N may rise many-fold
after a meal, and even urea-N excretion may increase modestly (reviewed by
Wood, 2001). These results
therefore support the prediction of Wood
(2001
) that elasmobranchs may
be so N-limited as to excrete virtually no `excess nitrogen' after feeding.
Inasmuch as ammonia-N excretion increased slightly, they also support the
prediction of Mommsen and Walsh
(1991
) that any elevation in
N-loss that does occur should be in the form of ammonia-N, because of the high
cost of urea-N synthesis.
This virtual absence of a post-prandial rise in N-excretion was not an
artifact of the experimental feeding approach or disturbance, because a very
similar result (no change in urea-N excretion, small rise in ammonia-N
excretion) was seen in our companion study on naturally feeding, uncannulated
dogfish (M.K., P.J.W., T.P.M. and C.M.W., unpublished results), where plasma
urea-N, ammonia-N and TMAO-N concentrations were similar to those of the
present study. Notably, in that study, total N-excretion rates were only
60% of those reported here, both before and after feeding, principally
due to a 40% lower rate of urea-N excretion. Despite the precautions taken to
minimise disturbance in Series 2, it appears that cannulation and experimental
handling inevitably increase the urea-N `leakiness' of the animals. Similarly
elevated baseline rates were reported in an earlier cannulation study on
S. acanthias (Wood et al.,
1995
) and possibly may result from an inhibitory action of
catecholamines on the active urea retention mechanism in the gills
(Pärt et al., 1998
),
discussed below.
The urea-dependent osmoregulatory strategy of elasmobranchs, as first
documented by Smith (1930), entails the maintenance of a massive urea-N
gradient from body fluids to seawater, and a substantial, continuous loss of
urea-N across the gills is unavoidable even under normal conditions
(Boylan, 1967;
Perlman and Goldstein, 1988
).
At the pre-feeding N-excretion rates (ammonia-N=60 µmol-N
kg1 h1, urea-N=750 µmol-N
kg1 h1) measured in the present study
(Figs 5,
6), the N-content of the meal
(32 500 µmol-N kg1 dogfish) would be lost in
40 h,
or in
67 h at the lower excretion rates measured (M.K., P.J.W., T.P.M.
and C.M.W., unpublished results) for uncannulated animals. These simple
calculations, coupled with an extensive literature indicating that feeding is
sporadic, irregular and opportunistic in sharks
(Jones and Geen, 1977
;
Cortes and Gruber, 1990
;
Hanchet, 1991
;
Tanasichuk et al., 1991
),
suggest that these elasmobranchs may be severely N-limited in nature, such
that N-conservation after feeding would be a high priority. Indeed, dietary
protein restriction eliminated the ability of the dogfish Scyliorhinus
stellaris to raise plasma urea-N in response to osmotic challenge
(Armour et al., 1993
). Plasma
urea-N levels and osmolality progressively fell during prolonged starvation in
S. acanthias (Leech et al.,
1979
) and in the pyjama shark, Poroderma africanum
(Haywood, 1973
), and rose
rapidly upon re-feeding in the latter. In the present study, plasma osmolality
increased significantly after feeding (Fig.
8A), but without any detectable post-prandial rises in plasma
urea-N (Fig. 7A). However, in
naturally feeding S. acanthias, osmolality and urea-N concentrations
in the blood plasma were significantly elevated at 20 h after a meal (M.K.,
P.J.W., T.P.M. and C.M.W., unpublished results); increases in plasma urea-N,
TMAO-N and Cl concentrations (discussed below) all
contributed to the phenomenon.
Several mechanisms probably contribute to N-conservation, particularly
after feeding. Firstly, the elasmobranch gill is uniquely designed to minimize
both urea-N and ammonia-N permeability, while maintaining permeability to
respiratory gases (Boylan,
1967). Wood et al.
(1995
) calculated that
branchial urea-N permeability in S. acanthias was only
7% of
that in a typical teleost, while branchial ammonia-N permeability was only
4%. A high cholesterol:phospholipid ratio in the basolateral membranes of
the gill epithelium appears to play an important role in this selective
permeability (Fines et al.,
2001
). Urea-N retention is also achieved by a low apical membrane
permeability and a `back-transport' mechanism for urea in the gill epithelium
(Pärt et al., 1998
) that
is basolaterally located, ATP-dependent and Na+-coupled
(Fines et al., 2001
). It would
be interesting to test whether the activity of this transporter increases
after feeding. Ammonia-N retention has been attributed to ammonia scavenging
by high-affinity glutamine synthetase in gill cells
(Wood et al., 1995
). However,
more importantly, glutamine synthetase activities increase substantially to
trap ammonia-N in a number of other tissues, particularly the liver, and there
is a clear activation of the ornithine urea cycle enzymes for urea-N synthesis
in both the liver and extra-hepatic tissues after natural feeding (M.K.,
P.J.W., T.P.M. and C.M.W., unpublished results). Finally, by analogy to
teleosts, it is likely that protein synthesis for growth is strongly
stimulated after feeding (reviewed by
Carter and Houlihan, 2001
;
Wood, 2001
), although to our
knowledge, this has never been evaluated in elasmobranchs. One indication that
this may be occurring is the decrease in
vO2 of venous blood
returning from the trunk musculature (Fig.
4), suggestive of increased metabolic expenditures in muscle at
this time.
Changes in plasma composition associated with feeding
Creatinine-N and TMAO-N were measured because of recent interest in the
metabolism of these N-products (see Walsh
and Mommsen, 2001) and the importance of the latter as a
stabilizing osmolyte (Yancey,
2001
). Creatinine is a breakdown product of muscle creatine, while
TMAO is probably of exogenous origin. Creatinine-N levels did not change after
experimental feeding (Table 2).
However, plasma TMAO-N levels rose very slightly
(Fig. 7C) both in this study
and in naturally feeding dogfish (M.K., P.J.W., T.P.M. and C.M.W., unpublished
results). Some elasmobranchs can synthesize TMAO, but it appears that S.
acanthias cannot (Goldstein et al,
1967
; Goldstein and Palatt,
1974
), so TMAO-N is thought to come from the diet, where it may be
more abundant than urea-N (e.g. Table
1).
In general, aerobic metabolism in elasmobranchs appears to rely heavily on
the oxidation of ketone bodies and, to a lesser extent, amino acids, with
reduced reliance on carbohydrate and almost no usage of fatty acids
(Ballantyne, 1997). Zammit and
Newsholme (1979) reported that prolonged starvation (up to 150 days) caused no
change in plasma glucose but a marked rise in plasma ß-hydroxybutyrate
levels in Sc. stellaris, while deRoos et al.
(1985
) described similar
responses in S. acanthias during a much shorter period of fasting (up
to 9 days). Richards et al.
(2003
) reported an increased
uptake of ß-hydroxybutyrate but no change in glucose uptake associated
with elevated post-exercise metabolism in a perfused trunk muscle preparation
of S. acanthias. The constancy of blood glucose and the modest
declines in ß-hydroxybutyrate and FAA-N levels in plasma after feeding in
the present study (Table 2) are
in accord with this scenario. Plasma FAA-N levels did not show an initial
surge after experimental feeding (Table
2), in marked contrast to teleosts, where large increases occur
(reviewed by Wood, 2001
). Only
a small delayed rise in plasma FAA-N levels at 30 h after natural feeding was
detected (M.K., P.J.W., T.P.M. and C.M.W., unpublished results). However,
plasma levels are not necessarily representative of flux. It is possible that
relatively slow digestion and absorption in the dogfish, coupled with
increased usage of amino acids as an oxidative substrate and for protein
synthesis, resulted in little apparent change in FAA-N concentrations.
Perhaps the most surprising change in plasma composition was the
progressive rise in Cl
(Fig. 8B), which explained
40% of the rise in plasma osmolality
(Fig. 8A). The same phenomenon
was seen in naturally feeding dogfish (M.K., P.J.W., T.P.M. and C.M.W.,
unpublished results). We hypothesise that the explanation lies in the powerful
acidbase exchange mechanisms in the gills of marine elasmobranchs
(Payan and Maetz, 1973
;
Bentley et al., 1976
;
Heisler, 1988
;
Claiborne and Evans, 1992
).
Apical Na+/H+ exchange powered by basolateral
Na+/K+-ATPase appears to occur in one type of
mitochondria-rich cell, and apical
Cl/HCO3 exchange powered by
basolateral V-H+-ATPase in another (Wilson et al.,
1997
,
2002
;
Piermarini and Evans, 2001
;
Piermarini et al., 2002
;
Claiborne et al., 2002
;
Edwards et al., 2002
;
Tresguerres et al., 2005
). An
ongoing activation of net Cl uptake in exchange for the
export of metabolic base by the latter mechanism i.e. compensation of
the alkaline tide would explain the increase in plasma
Cl. In future studies, it will be of interest to evaluate
whether the rectal gland (Shuttleworth,
1988
) is activated to handle this Cl load
indirectly associated with feeding.
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References |
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Andrade, D. V., De Toledo, L. F., Abe, A. S. and Wang, T.
(2004). Ventilatory compensation of the alkaline tide during
digestion in the snake, Boa constrictor. J. Exp. Biol.
207,1379
-1385.
Armour, K. J., O'Toole, L. B. and Hazon, N.
(1993). The effects of dietary protein restriction on the
secretory dynamics of 1-hydroxycorticosterone and urea in the dogfish,
Scyliorhinus canicula: a possible role for
1
-hydroxycorticosterone in sodium retention. J.
Endocrinol. 138,275
-282.[Abstract]
Babkin, B., Chaisson, A. and Friedman, M. (1935). Factors determining the course of the gastric secretion in elasmobranchs. J. Biol. Bd. Canada 1, 251-259.
Ballantyne, J. S. (1997). Jaws: the inside story. The metabolism of elasmobranch fishes. Comp. Biochem. Physiol. B 118,703 -742.[CrossRef]
Bentley, P. J., Maetz, J. and Payan, P. (1976). A study of the unidirectional fluxes of Na+ and Cl across the gills of the dogfish Scyliorhinus canicula (Chondrichthyes). J. Exp. Biol. 64,629 -637.[Abstract]
Boutilier, R. G., Heming, T. A. and Iwama, G. K. (1984). Appendix: physicochemical parameters for use in fish respiratory physiology. In Fish Physiology, Vol.10A (ed. W. S. Hoar and D. J. Randall), pp.403 -430. Orlando: Academic Press.
Boylan, J. W. (1967). Gill permeability in Squalus acanthias. In Sharks, Skates and Rays (ed. P. W. Gilbert, R. F. Mathewson and D. P. Rall), pp.197 -206. Baltimore: Johns Hopkins Press.
Bradford, M. (1976). A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72,248 -254.[CrossRef][Medline]
Caira, J. and Jolitz, E. (1989). Gut pH in the nurse shark, Ginglymostoma cirratum. Copeia 1, 192-194.
Cameron, J. N. (1971). Rapid method for
determination of total carbon dioxide in small blood samples. J.
Appl. Physiol. 31,632
-634.
Carter, C. G. and Houlihan, D. F. (2001). Protein synthesis. In Nitrogen Metabolism, Fish Physiology. Vol. 20 (ed. P. A. Anderson and P. A. Wright), pp. 31-75. Orlando: Academic Press.
Cho, C. Y. (1990). Fish nutrition, feeds, and feeding, with special emphasis on salmonid aquaculture. Food Rev. Int. 6,333 -357.
Claiborne, J. B. and Evans, D. H. (1992). Acidbase balance and ion transfers in the spiny dogfish (Squalus acanthias) during hypercapnia: a role for ammonia excretion. J. Exp. Zool. 261,9 -17.[CrossRef]
Claiborne. J. B., Edwards, S. L. and Morrison-Shetlar, A. I. (2002). Acidbase regulation in fishes: cellular and molecular mechanisms. J. Exp. Zool. 293,302 -319.[CrossRef][Medline]
Cortes, E. and Gruber, S. (1990). Diet, feeding habits, and estimates of daily ration of lemon sharks, Negaprion brevirostris. Copeia 1,204 -208.
DeBoeck, G., Grosell, M. and Wood, C. M. (2001). Sensitivity of the spiny dogfish (Squalus acanthias) to waterborne silver exposure. Aquat. Toxicol. 54,261 -275.[CrossRef][Medline]
deRoos, R., deRoos, C. C., Werner, C. S. and Werner, H. (1985). Plasma levels of glucose, alanine, lactate, and ß-hydroxybutyrate in the unfed spiny dogfish shark (Squalus acanthias) after surgery and following mammalian insulin infusion. Gen. Comp. Endocrinol. 58, 28-43.[CrossRef][Medline]
Edwards, S. L., Donald, J. A., Toop, T., Donowitz, M. and Tse, C. M. (2002). Immunolocalisation of sodium/proton exchanger-like proteins in the gills of elasmobranchs. Comp. Biochem. Physiol. A 131,257 -265.
Fines, G. A., Ballantyne, J. S. and Wright, P. A. (2001). Active urea transport and an unusual basolateral membrane composition in the gills of a marine elasmobranch. Am. J. Physiol. 280,R16 -R24.
Gilmour, K. M., Henry, R. P., Wood, C. M. and Perry, S. F.
(1997). Extracellular carbonic anhydrase and an acidbase
disequilibrium in the blood of the dogfish Squalus acanthias. J.
Exp. Biol. 200,173
-183.
Gilmour, K. M., Perry, S. F., Bernier, N. J., Henry, R. P. and Wood, C. M. (2001). Extracellular carbonic anhydrase in the dogfish, Squalus acanthias: A role in CO2 excretion. Physiol. Biochem. Zool. 74,477 -492.[CrossRef][Medline]
Goldstein, L. and Palatt, P. J. (1974).
Trimethylamine oxide excretion rates in elasmobranchs. Am. J.
Physiol. 227,1268
-1272.
Goldstein, L., Hartman, S. C. and Forster, R. P. (1967). On the origin of trimethylamine oxide in the spiny dogfish, Squalus acanthias. Comp. Biochem. Physiol. 21,719 -722.[CrossRef][Medline]
Hanchet, S. (1991). Diet of spiny dogfish, Squalus acanthias, on the east coast, South Island, New Zealand. J. Fish. Biol. 39,313 -323.
Haywood, G. P. (1973). Hypo-osmotic regulation coupled with reduced metabolic urea in the dogfish Poroderma africanum: an analysis of serum osmolarity, chloride, and urea, Mar. Biol. 23,121 -127.[CrossRef]
Heinegarde, D. and Tiderstrom, G. (1973). Determination of serum creatinine by a direct colorimetric method. Clin. Chim. Acta 43,305 .[CrossRef][Medline]
Heisler, N. (1988). Acidbase regulation. In Physiology of Elasmobranch Fishes (ed. T. J. Shuttleworth), pp. 215-252. Berlin: Springer-Verlag.
Henry, R. P., Gilmour, K. M., Wood, C. M. and Perry, S. F. (1997). Extracellular carbonic anhydrase activity and carbonic anhydrase inhibitors in the circulatory system of fish. Physiol. Zool. 70,650 -659.[Medline]
Hersey, S. J. and Sachs, G. (1995).
Gastric-acid secretion. Physiol. Rev.
75,155
-189.
Jones, B. C. and Geen, G. H. (1977). Food and feeding of spiny dogfish (Squalus acanthias) in British Columbia waters. J. Fish. Res. Bd. Canada 34,2067 -2078.
Kajimura, M., Croke, S. J., Glover, C. N. and Wood, C. M. (2004). Dogmas and controversies in the handling of nitrogenous wastes; The effect of feeding and fasting on the excretion of ammonia, urea, and other nitrogenous waste products in rainbow trout. J. Exp. Biol. 15,1993 -2002.[CrossRef]
Leech, A. R., Goldstein, L., Cha, C. J. and Goldstein, J. M. (1979). Alanine biosynthesis during starvation in skeletal muscle of the spiny dogfish, Squalus acanthias. J. Exp. Zool. 207, 73-80.[CrossRef]
Lenfant, C and Johansen, K. (1966). Respiratory function in the elasmobranch Squalus suckleyi G. Respir. Physiol. 1,13 -29.[CrossRef][Medline]
Maren, T. H. (1967). Carbonic anhydrase:
Chemistry, physiology, and inhibition. Physiol. Rev.
47,595
-781.
McMurray, C. H., Blanchflower, W. J. and Rice, D. A.
(1984). Automated kinetic method for D-3-hydroxybutyrate in
plasma or serum. Clin. Chem.
30,421
-425.
Menon, M. and Kewalramani, H. (1959). Studies on some physiological aspects of digestion in three species of elasmobranchs. Proc. Ind. Acad. Sci. B 50, 26-39.
Mommsen, T. P. and Walsh, P. J. (1991). Urea synthesis in fishes: evolutionary and biochemical perspectives. In Biochemistry and Molecular Biology of Fishes. Vol.1 (ed. P. W. Hochachka and T. P. Mommsen), pp.137 -163. New York: Elsevier.
Mondzac, A., Ehrlich, G. E. and Seegmiller, J. E. (1965). An enzymatic determination of ammonia in biological fluids. J. Lab. Clin. Med. 66,526 -531.[Medline]
Moore, S. (1968). Amino acid analysis: aqueous
dimethyl sulfoxide as solvent for the ninhydrin reaction. J. Biol.
Chem. 243,6281
-6283.
Niv, Y. and Fraser, G. M. (2002). The alkaline tide phenomenon. J. Clin. Gastroenterol. 35, 5-8.[CrossRef][Medline]
Papastamatiou, Y. P. and Lowe, C. G. (2004).
Postprandial response of gastric pH in leopard sharks (Triakis
semifasciata) and its use to study foraging ecology. J. Exp.
Biol. 207,225
-232.
Pärt, P., Wright, P. A. and Wood, C. M. (1998). Urea and water permeability in dogfish gills (Squalus acanthias). Comp. Biochem. Physiol. A 119,117 -123.
Payan, P. and Maetz, J. (1973). Branchial sodium transport mechanisms in Scyliorhinus canicula: evidence for Na+/NH4+ and Na+/H+ exchanges and for a role of carbonic anhydrase. J. Exp. Biol. 58,487 -502.
Perry, S. F. and Wood, C. M. (1989). Control and co-ordination of gas transfer in fishes. Can. J. Zool. 67,2961 -2970.
Perlman, D. F. and Goldstein, L. (1988). Nitrogen metabolism. In Physiology of Elasmobranch Fishes (ed T. J. Shuttleworth), pp.253 -275. Berlin: Springer-Verlag.
Piermarini, P. M. and Evans, D. H. (2001).
Immunochemical analysis of the vacuolar proton-ATPase B-subunit in the gills
of a euryhaline stingray (Dasyatis sabina): effects of salinity and
relation to Na+/K+-ATPase. J. Exp.
Biol. 204,3251
-3259.
Piermarini, P. M., Verlander, J. W., Royaux, I. E. and Evans, D. H. (2002). Pendrin immunoreactivity in the gill epithelium of a euryhaline elasmobranch Am. J. Physiol. 283,R983 -R992.
Rahmatullah, M. and Boyde, T. R. (1980). Improvements in the determination of urea using diacetyl monoxime: methods with and without deproteinization. Clin. Chem. Acta 107, 3-9.[CrossRef][Medline]
Richards, J. G., Heigenhauser, G. F. and Wood, C. M. (2003). Exercise and recovery metabolism in the Pacific spiny dogfish (Squalus acanthias). J. Comp. Physiol. B. 173,463 -474.[Medline]
Rune, S. J. (1965). The metabolic alkalosis following aspiration of gastric acid secretion. Scand. J. Clin. Lab. Invest. 17,305 -310.[Medline]
Rune, S. J. (1966). Comparison of the rates of gastric acid secretion in man after ingestion of food and after maximal stimulation with histamine. Gut 7, 344-350.[Medline]
Sachs, G., Sims, J. M., Briving, C., Wallmark, B. and Hersey, S. (1995). The pharmacology of the gastric acid pump: the H+, K+-ATPase. Annu. Rev. Pharmacol. Toxicol. 35,277 -305.[CrossRef][Medline]
Schurdak, M. and Gruber, S. (1989). Gastric evacuation of the lemon shark Negaprion brevirostris under controlled conditions. Exp. Biol. 48, 77-82.[Medline]
Shuttleworth, T. J. (1988). Salt and water balance extrarenal mechanisms. In Physiology of Elasmobranch Fishes (ed. T. J. Shuttleworth), pp.171 -199. Berlin: Springer-Verlag.
Sims, D., Davies, S. and Bone, Q. (1996). Gastric emptying rate and return of appetite in lesser spotted dogfish, Scyliorhinus canicula. J. Mar. Biol. Assoc. UK 76,476 -491.
Smith, H. W. (1936). The retention and physiological role of urea in the elasmobranchii. Biol. Rev. 11,49 -82.
Smolka, A., Lacy, E., Luciano, L. and Reale, E.
(1994). Identification of gastric H,K ATPase in an early
vertebrate, the Atlantc stingray Dasyatis sabina. J. Histochem.
Cytochem. 42,1323
-1342.
Sullivan, M. (1905). The physiology of the
digestive tract of elasmobranchs. Am. J. Physiol.
15, 42-45.
Tanasichuk, R. W., Ware, D. M., Shaw, W. and McFarlane, G. A. (1991). Variations in diet, daily ration, and feeding periodicity of Pacific hake (Merluccius productus) and spiny dogfish (Squalus acanthias) off the lower coast of Vancouver Island. Can. J. Fish. Aquat. Sci. 48,2118 -2128.
Tresguerres, M., Katoh, F., Fenton, H. and Goss, G.
(2005). Regulation of branchial V-H+-ATPase,
Na+/K+-ATPase and NHE2 in response to acid and base
infusions in the Pacific spiny dogfish (Squalus acanthias).
J. Exp. Biol. 208,345
-354.
Verdouw, H., van Echted, C. J. A. and Dekkers, E. M. J. (1978). Ammonia determination based on indophenol formation with sodium salicylate. Water. Res. 12,399 -402.[CrossRef]
Walsh, P. J. and Mommsen, T. P. (2001). Evolutionary considerations of nitrogen metabolism and excretion. In Nitrogen Metabolism, Fish Physiology. Vol.20 (ed. P. A. Anderson and P. A. Wright), pp.1 -30. Orlando: Academic Press.
Wang, T., Busk, M. and Overgaard, J. (2001). The respiratory consequences of feeding in amphibians and reptiles. Comp. Biochem. Physiol. A 128,535 -549.
Wekell, J. C. and Barnett, H. (1991). New method for analysis of trimethylamine oxide using ferrous sulfate and EDTA. J. Food Sci. 56,132 -138.
Wetherbee, B. M. and Gruber, S. H. (1990). The effects of ration level on food retention time in juvenile lemon sharks, Negaprion brevirostris. Environ. Biol. Fish. 29, 59-65.[CrossRef]
Wetherbee, B. M., Gruber, S. H. and Ramsay, A. L. (1987). X-radiographic observation of food passage through digestive tracts of lemon sharks. Trans. Am. Fish. Soc. 116,763 -767.[CrossRef]
Williams, H., McVicar, A. and Ralph, R. (1970). The alimentary tact of fish as an environment for helminth parasites. In Aspects of Fish Parasitology (ed. A. Taylor and R. Muller), pp. 43-79. Oxford: Blackwell Scientific Publications.
Wilson, J. M., Randall, D. J., Vogl, A. W., Harris, J. and Iwama, G. (1997). Immunolocalization of proton-ATPase in the gills of the elasmobranch, Squalus acanthias. J. Exp. Zool. 278,78 -86.[CrossRef][Medline]
Wilson, J. M., Randall, D. J., Vogl, A. W., Harris, J., Sly, W. S. and Iwama, G. K. (2000). Branchial carbonic anhydrase is present in the dogfish, Squalus acanthias. Fish Physiol. Biochem. 22,329 -336.[CrossRef]
Wilson, J. M., Morgan, J. D., Vogl, A. W. and Randall, D. J. (2002). Branchial mitochondria-rich cells in the dogfish Squalus acanthias. Comp. Biochem. Physiol. A 132,365 -374.
Wood, C. M. (2001). The influence of feeding, exercise, and temperature on nitrogen metabolism and excretion. In Nitrogen Metabolism, Fish Physiology. Vol.20 (ed. P. A. Anderson and P. A. Wright), pp.201 -238. Orlando: Academic Press.
Wood, C. M., Perry, S. F., Walsh, P. J. and Thomas, S. (1994). HCO3 dehydration by the blood of an elasmobranch in the absence of a Haldane effect. Respir. Physiol. 98,319 -337.[CrossRef][Medline]
Wood, C. M., Pärt, P. and Wright, P. A.
(1995). Ammonia and urea metabolism in relation to gill function
and acidbase balance in a marine elasmobranch, the spiny dogfish
(Squalus acanthias). J. Exp. Biol.
198,1545
-1558.
Yancey, P. H. (2001). Nitrogen compounds as osmolytes. In Nitrogen Metabolism, Fish Physiology, vol. 20 (ed. P. A. Anderson and P. A. Wright), pp.309 -341. Orlando: Academic Press.
Zammitt, V. A. and Newsholme, E. A. (1979). Activities of enzymes of fat and ketone-body metabolism and effects of starvation on blood concentration of glucose and fat fuels in teleost and elasmobranch fish. Biochem. J. 184,313 -322.[Medline]
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