All rainbow trout (Oncorhynchus mykiss) are not created equal: intra-specific variation in cardiac hypoxia tolerance
1 Department of Biology, Portland State University, PO Box 0751, Portland,
OR 97207-0751, USA
2 Department of Biological Sciences, Idaho State University, Pocatello, ID
82309-8007, USA
Author for correspondence at present address: Ocean Sciences Center, Memorial
University of Newfoundland, St John's, Newfoundland, Canada. A1C 5S7 (e-mail:
kgamperl{at}mun.ca)
Accepted 8 December 2003
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Summary |
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Key words: rainbow trout, Oncorhynchus mykiss, intra-specific variation, cardiac hypoxia tolerance, heart, lactate dehydrogenase
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Introduction |
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The rainbow trout is generally considered to be a hypoxia-sensitive species
(Gesser, 1977;
Dunn and Hochachka, 1986
;
Arthur et al., 1992
;
Gamperl et al., 2001
).
However, we have identified an aquaculture facility in Oregon (USA) that
produces rainbow trout which appear to have a considerable degree of inherent
myocardial hypoxia tolerance. For example, although Gamperl et al.
(2001
) report that 15 min of
severe hypoxia (PO <5 mmHg; 1 mmHg=133.3 Pa) with only
5 min of physiological afterload (50 cmH2O; 1
cmH2O=98.07 Pa) reduced post-hypoxic cardiac output in trout from a
British Columbia (Canada) facility by 38%, this protocol had no effect on
post-hypoxic cardiac function in these Oregon-reared trout. In the current
study, we determine the degree of myocardial hypoxia-tolerance displayed by
these rainbow trout, and investigate whether myoglobin or lactate
dehydrogenase release from the myocardium can be used as indices of myocardial
damage (necrosis) in the in situ perfused trout heart. This latter
goal is important for future studies of myocardial hypoxia tolerance and
preconditioning in fishes. The loss of cardiac function following severe
hypoxia could occur in response to temporary contractile dysfunction (i.e.
stunning; Ferrari et al.,
1999
) or permanent cellular necrosis, and the quantification of
cell death remains the most widely accepted end-point for identifying
myocardial preconditioning (Wolff et al.,
2000
).
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Materials and methods |
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Surgical procedures
All procedures were approved by the Animal Care Committee at PSU, and
conformed with the Guide for the Care and Use of Laboratory Animals published
by the US National Institutes of Health (NIH Publication No. 85-23, revised
1996). Trout were anesthetized in an oxygenated, buffered solution of tricaine
methane sulfonate (0.1 g l1 MS-222; 0.1 g
l1 sodium bicarbonate) and transferred to an operating table
where their gills were irrigated with oxygenated, buffered anesthetic (0.05 g
l1 MS-222; 0.05 g l1 sodium bicarbonate)
at 46°C. Fish were then injected with 1.0 ml of heparinized (100
i.u. ml1) saline via the caudal vessels, and an
in situ heart preparation was obtained as detailed in Farrell et al.
(1986).
The saline used to perfuse the heart (pH 7.8 at 10°C) contained (in
mmol l1): NaCl (124), KCl (3.1),
MgSO4.7H2O (0.93), CaCl2.2H2O
(2.52), glucose (5.0), TES salt (6.4) and TES acid (3.6)
(Keen et al., 1993). These
chemicals were purchased from Fisher Scientific (Fair Lawn, NJ, USA), with the
exception of the TES salt (Sigma Chemical Co., St Louis, MO, USA). Adrenaline
bitartrate (15 nmol l1 final concentration; Sigma Chemical
Co.) was added to the perfusate every 20 min throughout the experiment to
ensure the long-term viability of the perfused trout heart
(Graham and Farrell, 1989
).
The saline was bubbled with 100% O2 for a minimum of 45 min prior
to use. Although the coronary circulation was not perfused, prior research on
fish of similar size suggests that this level of oxygenation can supply
sufficient O2 to the outer myocardium such that the maximum
performance of the in situ heart is comparable
(Farrell et al., 1986
) and
perhaps even higher (Farrell et al.,
1991
) than that measured in vivo. For hypoxic exposures,
the perfusate was bubbled with 100% N2 for a minimum of 2 h prior
to the experiments to ensure that PO2 was
510 mmHg. Potential oxygen transfer from the experimental bath to the
heart was minimized by covering the bath with a loose-fitting plastic lid, and
by bubbling 100% N2 into the bath beginning 5 min prior to the
onset of severe hypoxia.
Experiment 1
Before assessing the degree of myocardial hypoxia tolerance displayed by
these trout hearts, we wanted to ensure that the hearts were truly exposed to
severely hypoxic conditions when perfused with N2-equilibrated
saline in our experimental apparatus.
Two groups (N=78) of in situ hearts were exposed
to 15 min of severe hypoxia, at an output pressure (Pout)
of 50 cmH2O (Fig.
1). In addition, one of the groups was also exposed to 1.5 mmol
l1 of NaCN. Cardiac output
() and heart rate (fH) were
monitored at 2 min intervals throughout severe hypoxia. In these experiments,
the recovery of maximum cardiac output
(
MAX) was not assessed, due
to the potential effects of residual NaCN on in situ cardiac
performance. In this experiment, the PO2 of the
perfusate entering the heart was also measured by collecting perfusate samples
(12 ml) in gas tight Hamilton syringes and injecting each sample into a
water-jacketed E-101 oxygen electrode at 10°C (Cameron Instrument Company,
Port Aransas, TX, USA). PO2 (in mmHg) was read
from an OM-200 dual channel oxygen meter (Cameron Instrument Company, Port
Aranas, TX, USA).
|
Experiment 2
This experiment assessed the degree of myocardial hypoxia tolerance
displayed by trout from Clear Creek Rainbow Ranch. In this experiment, each
protocol was separated into 3 main sections: (1) stabilization and
MAX1, (2) the experimental
period and (3) recovery and
MAX2. All cardiovascular
variables (input pressure, PIN; output pressure,
POUT; and
) were
manipulated in an identical manner during the initial and final portions of
each protocol. However, the protocols were unique in terms of the duration of
severe hypoxia administered during the experimental period.
Stabilization and MAX1
Once the fish was placed into the experimental bath and connected to the
perfusion apparatus, PIN was set to achieve a
physiologically relevant (1617
ml min1 kg1;
Kiceniuk and Jones, 1977
), and
POUT was maintained at 10 cmH2O for 5 min.
Thereafter, POUT was raised to 50 cmH2O, a
level comparable to in vivo arterial pressures
(Kiceniuk and Jones, 1977
).
After allowing the heart to stabilize at a POUT of 50
cmH2O for 5 min, PIN was gradually increased
until
reached 30 ml
min1 kg1. This initial cardiac stretch,
which was maintained for 20 s, allowed any air bubbles to be cleared from
within the heart and provided an initial assessment of cardiac viability.
Hearts were discarded if they required more than a 3 cmH2O increase
in PIN to reach a
of 30 ml min1 kg1, and were assumed to
have either poor cannula placement, cannula obstruction or myocardial
damage.
Following the cardiac stretch, all hearts were maintained at a
of 1617 ml
min1 kg1 for 20 min before their initial
maximum cardiac output
(
MAX1) was determined.
Maximum cardiac output
(
MAX) was achieved by
increasing PIN in a stepwise fashion from that required to
achieve resting cardiac output (
1.0 cmH2O) to 3.0
cmH2O, to 4.0 cmH2O, and finally to 4.5 cmH2O
(Fig. 2). Each stepwise
increase in PIN was maintained for approximately 20 s, and
resting
was quickly re-established
after
MAX was reached. The
entire
MAX test took
approx. 5 min to complete. After
MAX1 had been measured,
hearts were randomly assigned to a treatment group.
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Experimental period
Hearts were exposed to either (A) a control treatment (oxygenated
perfusion) (N=7), or to severe hypoxia for (B) 10 min,
(N=7), (C) 20 min (N=7) or (D) 30 min (N=8)
(Fig. 2). To ensure that the
total length of each treatment was equal, despite the variable durations of
hypoxic exposure, the period of resting (oxygenated) cardiac function
preceding hypoxia ranged from 35 to 65 min.
Throughout the experimental period, POUT was set at 50
cmH2O. During all periods of oxygenated perfusion
was maintained at a resting level of
1617 ml min1 kg1 by adjusting
PIN. However, PIN was not increased to
maintain
during severe hypoxia
because several preliminary experiments showed that the in situ
hearts failed to regain contractile function when an attempt was made to
maintain pre-hypoxic workloads (data not shown).
Recovery and MAX2
Immediately following the main hypoxic period, the in situ heart
was perfused with oxygenated saline, and a resting
of 1617 ml
min1 kg1 was quickly restored (within
24 min). This was accomplished by setting POUT at a
sub-physiological level (10 cmH2O) and gradually increasing
PIN. Lowering POUT to 10 cm
H2O facilitated the rapid recovery of cardiac function following
hypoxia by reducing the work required by the heart to generate a given flow.
Following this 10 min period of reduced after-load, POUT
was restored to 50 cmH2O and the heart was allowed to recover for
20 min before the final maximum cardiac output test
(
MAX2) was administered
(Fig. 2). This test was
performed using the same procedures described for the
MAX1 test.
After had been restored to
1617 ml min1 kg1 following
MAX2, two tests were
performed to ensure that the input cannula was securely tied into the sinus
venosus, and that the in situ heart was isolated from the saline in
the experimental bath. First, it was confirmed that
rapidly fell to 0 after the input
cannula was clamped off with a pair of haemostats. Second, with the haemostats
still clamping the input cannula, and the tubing connected to the output
cannula raised to 100 cmH2O, we ensured that there was no backflow
of perfusate. After completing these tests, the ventricle was rapidly excised,
blotted to remove residual saline, and weighed.
Perfusate samples (1 ml) for protein and myoglobin analyses were collected immediately prior to the main hypoxic challenge, and at 2, 4, 6, 10, 15, 20 and 30 min following hypoxia (Fig. 2). During the control treatments, perfusate samples were taken at points equivalent to those used in the other treatment groups. The perfusate and ventricular samples were immediately frozen in liquid nitrogen, and stored at 70°C for subsequent biochemical analysis.
Data collection and analysis
Cardiac function was continuously monitored throughout each experiment by
measuring , PIN and
POUT. Cardiac output (ml min1) was
measured using a Model T206 small animal blood flow meter in conjunction with
a pre-calibrated in-line flow probe (2 N, Transonic Systems Inc., Ithaca, NY,
USA). Gould Statham pressure transducers (P23 ID, Oxnard, CA, USA) were used
to measure PIN and POUT
(cmH2O). The pressure transducers were calibrated daily against a
static column of water, where zero pressure (0 cmH2O) was set equal
to the saline level in the experimental bath. In addition, the recorded input
and output pressures were corrected to account for the resistance in the
tubing between the points of pressure measurement and the heart, using
predetermined calibrations.
Signals from the flow meter and the pressure transducers were amplified and
filtered using a Model MP100A-CE data acquisition system (BIOPAC Systems Inc.,
Santa Barbara, CA, USA). The acquired signals were then analyzed and stored
using Acqknowledge Software (BIOPAC Systems). Although data were continuously
collected, cardiovascular function was only analyzed at specific intervals
during each experiment. The resting PIN required to
maintain a of 1617 ml
min1 kg1 was measured prior to the
MAX1 and
MAX2 tests. A rise in
resting PIN over the course of the experiments was used as
an index of diminished resting cardiac function. Because
MAX tests were administered
at the beginning and at the end of each experiment (Figs
2 and
3), reductions in
MAX and maximum stoke
volume (VS) were used as measures of reduced maximum heart function.
Heart rate (fH) and
were
also measured at regular intervals (every 25 min) throughout the
hypoxic period to provide an index of cardiac function during hypoxia.
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Heart rate was calculated by measuring the number of systolic peaks during
a 2030 s interval and stroke volume VS was calculated as
/fH.
Biochemical assessment of myocardial damage
To determine when myoglobin concentrations in the perfusate would probably
be at their maximum, the total concentration of protein in the perfusate was
quantified between 2 and 30 min post-hypoxia using the Bradford dye-binding
procedure (Bradford, 1976),
with bovine serum albumin (BSA) as the standard. No protein was detected in
the perfusate leaving in situ hearts exposed to the control treatment
or to 10 min of severe hypoxia. However, protein was detected in two-thirds of
the hearts exposed to 20 and 30 min of severe hypoxia at 2, 4 or 6 min of
recovery (range 0.310 µg ml1).
Myoglobin
Based on the results of the protein assay, the amount of myoglobin released
from the heart was assessed in perfusate samples collected 4 min following
hypoxia. Samples (20 µl) were solubilized in Laemmli sample buffer
containing 2% SDS and dithiothreitol (final concentration 1 mmol
l1) at 70°C for 10 min. In addition, a frozen tissue
standard of trout cardiac muscle (approx. 50 mg) was homogenized in 19 volumes
of filtered (0.22 µm), ice-cold extraction buffer (20 mmol
l1 Hepes, 250 mmol l1 sucrose, 1 mmol
l1 EDTA, pH 7.5). This homogenate was then centrifuged
(600 g) for 10 min at 4°C, and the supernatant was used
for myoglobin measurements.
All samples were electrophoresed (BioRad Mini Protean II, Hercules, CA, USA) at 140 mV for 2 h with a 17.5% TricineSDS polyacrylamide resolving gel. Immediately after electrophoresis, proteins were transferred to PVDF membranes (0.45 µm; Immobilon-P, Millipore Corp., Bedford, MA, USA) using a Mini Transblot apparatus (BioRad) set at 150 mA for 50 min. These membranes were then soaked overnight at 4°C in phosphate-buffered saline (PBS) (pH 7.4) containing 5% nonfat dry milk (Carnation, Los Angeles, CA, USA). Immunoblotting was performed with a polyclonal rabbit anti-myoglogin antibody (Sigma #M8648; diluted 1:1000 in PBS containing 0.1% BSA and 0.02% sodium azide) for 60 min at 25°C. Membranes were then washed in PBS containing 1% Triton X-100, and incubated for 60 min at 25°C with goat anti-rabbit HRP-conjugated IgG (BioRad) that was diluted 1:15 000 in PBS containing 0.1% BSA without sodium azide. Enhanced chemiluminescence (Amersham Life Sciences, Buckinghamshire, UK) was used to visualize bands using the Fluor-S Multi-imaging system (BioRad). The relative amount of protein in each band was quantified using scanning densitometry and Quantity One software (BioRad).
Lactate dehydrogenase
Excessive dilution caused by high perfusate flow rates (1617 ml
min1 kg1) and the absence of perfusate
recirculation may prevent the detection of metabolic enzymes in the perfusate
leaving in situ hearts (Gamperl
et al., 2001). Therefore, LDH activity remaining in the ventricle
was quantified following each of the treatments.
In addition, LDH activity was quantified in ventricles from a group of baseline (non-experimental) trout (N=6) that were sampled directly from the holding tanks. Baseline trout were euthanized using cerebral percussion, then the hearts were quickly (<30 s) excised and allowed to beat in ice-cold saline for 1 min to clear any residual blood from within the heart. Ventricular LDH activities in the control and baseline hearts were compared to determine whether any myocardial cell death occurred as a result of surgery and/or the duration of the experimental protocol. Further, the LDH activity in the baseline ventricles was compared to literature values in order to evaluate whether elevated ventricular LDH activities could explain the enhanced hypoxia tolerance observed in this population of rainbow trout.
Samples of frozen ventricle (50100 mg) were homogenized in 9 volumes of ice-cold extraction buffer (50 mmol l1 Hepes, 1 mmol l1 EDTA, and 2 mmol l1 dithiothreitol, pH 7.6 at 10°C) using a motorized Duall-21 ground-glass homogenizer. The reaction medium containing whole homogenate, 50 mmol l1 Hepes, 1 mmol l1 KCN, 0.17 mmol l1 NADH and 1 mmol l1 pyruvate (omitted from controls), was maintained at pH 7.6 and 10°C. LDH activity was analyzed using a Perkin-Elmer Lambda 6 UV/VIS spectrophotometer (Norwalk, CT, USA) equipped with a thermostatically controlled recirculating water bath and water-jacketed cuvette holder. The reaction was followed at 340 nm for 5 min. The analytical grade biochemicals used in this analysis were purchased from Sigma Chemical Co.
Statistics
All statistical analyses were performed using StatView Software (SAS
Institute Inc., Cary, NC, USA). One-way analyses of variance (ANOVAs),
followed by Fisher's protected least significant difference (PLSD)
post-hoc tests, were used to compare parameters between the treatment
groups, including: (1) body and ventricular mass, (2) resting cardiac function
(, VS and fH) prior to
MAX1, (3) maximum cardiac
function (
, VS and fH) at
MAX1, (4) the percent
change in maximum cardiac performance
(
MAX2 versus
MAX1), (5) the percent change
in resting PIN prior to
MAX1 versus
MAX2 and (6) average
ventricular LDH activities. Repeated-measures ANOVAs were performed for all
comparisons of (1) maximum myocardial performance
(
MAX1 versus
MAX2) within each treatment
group, (2) resting PIN (prior to
MAX2 versus prior
to
MAX1) within each
treatment group and (3) the loss of cardiac function
(
and fH) during 30 min of
severe hypoxia between the treatment groups. The level of statistical
significance used in each analysis was P<0.05. All percentage data
were arc-sine transformed prior to running any statistical tests.
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Results |
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Experiment 2
Ventricular mass (0.52±0.02 g) and relative ventricular mass
(0.09±0.003%) were not significantly different between the various
treatments. Therefore, and
VS are reported on a mass-specific basis (ml min1
kg1 body mass).
Initial cardiac function under oxygenated conditions
Prior to MAX1, resting
cardiovascular function was not significantly different between treatments.
When the data for all groups was combined, a PIN of
1.0±0.4 cmH2O was required to maintain a resting
of 17.0±0.2 ml
min1 kg1, and resting fH and
VS values were 69.3±3.7 beats min1 and
0.26±0.02 ml kg1, respectively. Furthermore, there
were no significant differences in
(54.4±3.1 ml min1 kg1), VS
(0.95±0.05 ml kg1) or fH (58.4±3.2
beats min1) at
MAX1.
Cardiac function during severe hypoxia
Cardiac performance was compromised to varying degrees during severe
hypoxia. Some in situ hearts generated positive flow throughout the
hypoxic challenge, while fell to zero
in others. In addition, many in situ hearts developed an irregular
fH during severe hypoxia, and/or during initial recovery from the
hypoxic challenge. Overall cardiovascular function decreased at a similar rate
in hearts exposed to 10, 20 or 30 min of severe hypoxia
(Fig. 4). However, in all
treatments,
(Fig. 4A) decreased more
rapidly than fH (Fig.
4B). Cardiac output decreased by 66% (falling from 16.8±0.4
ml min1 kg1 to 5.7±1.2 ml
min1 kg1) during the first 10 min of
severe hypoxia, and ultimately reached 2.4±0.9 ml
min1 kg1 (approx. 15% of the initial
value) after 30 min of severe hypoxia (Fig.
4). In contrast, fH slowed by approx. 20% every 10 min,
and reached 30.5±7.1 beats min1 (approx. 40% of the
initial value) by 30 min of severe hypoxia
(Fig. 4B). Because
fell more dramatically and more
rapidly than fH, it is clear that VS was also reduced (by
approx. 64%) during the 30 min hypoxic challenge (data not shown).
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Cardiac function following reperfusion
A significantly greater PIN (as compared with
pre-hypoxic values) was required to maintain a resting
of 1617 ml
min1 kg1 following 10 min (0.8±0.1
cmH2O) and 30 min (1.1±0.3 cmH2O) of severe
hypoxia (Fig. 5). Maximum
fH also increased following the control (by 7.0±1.8 beats
min1) and 20 min (by 5.5±1.9 beats
min1) of severe hypoxia treatments
(Fig. 6). However, these
increases in resting PIN and maximum fH were not
significantly different when all groups were compared (Figs
5B and
6B, respectively). Therefore,
the effect of the duration of severe hypoxia on cardiac function was evaluated
by comparing changes in maximum VS and
MAX.
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|
Maximum VS and
MAX were not affected by
the control protocol. However, a strong negative relationship existed between
the duration of severe hypoxia and the recovery of maximum cardiac function
(Fig. 6B). Maximum
decreased by 4.2±2.2%,
15.3±5.6% and 23.0±5.0% after hearts were exposed to 10, 20 or
30 min of severe hypoxia, respectively. Further, although the reduction in
VS following 10 min of severe hypoxia was not significantly greater
as compared with the control treatment, significant decreases in VS
were recorded after both 20 min (23.2±5.5%) and 30 min
(27.4±6.5%) of severe hypoxia.
Biochemical markers of myocardial damage
Myoglobin was not detected in the perfusate 4 min following severe hypoxia,
even though the trout heart standard was clearly labeled by the polyclonal
rabbit anti-myoglobin antibody (data not shown). For baseline
(non-experimental) fish, ventricular LDH activity was 131.1±7.6 units
g1 wet mass (1 unit of LDH activity = conversion of 1
µmol substrate to product per min), a value not significantly different
from that measured in the control group (172.3±21.9 units
g1 wet mass) (Fig.
7). Furthermore, ventricular LDH activity did not vary
significantly in response to the duration of hypoxic exposure
(Fig. 7). These results suggest
that myocardial necrosis did not occur in response to severe hypoxia.
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Discussion |
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Trout cardiac function and hypoxia tolerance
The in situ hearts in this study experienced an 85% drop in
resting , a 60% fall in fH,
and a 65% decrease in VS (data not shown) during 30 min of severe
hypoxia (PO =510 mmHg)
(Fig. 4). This magnitude of
functional loss is very similar to that reported in previous studies of
hypoxic cardiac function using in situ
(Gamperl et al., 2001
) or
in vitro (Gesser,
1977
) trout heart preparations. These similarities, in hypoxic
myocardial function, when combined with the data from Experiment 1, strongly
suggest that the in situ hearts in this study were indeed exposed to
severe hypoxia and showed a typical functional response to oxygen
deprivation.
Although 30 min of severe hypoxia significantly reduced maximum cardiac
function in Experiment 2, several pieces of evidence indicate that these
in situ trout hearts were extremely hypoxia-tolerant, as compared
with other rainbow trout. Gamperl et al.
(2001) found that in
situ rainbow trout hearts at Simon Fraser University experienced a
significant decline in
MAX
(23%) following only 15 min of severe hypoxia with POUT at
10 cmH2O. However, the identical protocol to that used by Gamperl
et al. (2001
) did not
significantly affect
MAX in
the present study, and a comparable loss of function (23% loss of
MAX) could only be achieved
by doubling the duration of hypoxia (from 15 to 30 min), and by increasing the
workload of the hearts during the hypoxic challenge fivefold (i.e. by making
the hearts pump against a physiological POUT of 50
cmH2O) (Fig. 2).
Furthermore, the degree of functional recovery in these in situ trout
hearts (approx. 77%) following 30 min of severe hypoxia was much closer to
that measured in ventricular strips from the carp (approx. 90%) than from the
rainbow trout (approx. 40%; Gesser,
1977
) (Fig. 8). The
carp is a species capable of surviving up to 4.5 months of environmental
hypoxia (Piironen and Holopainen,
1986
). Although inter-specific differences in myocardial hypoxia
tolerance are expected (Driedzic and
Gesser, 1994
), these data strongly suggest that significant
intra-specific variation in myocardial hypoxia tolerance exists in fishes.
|
Possible factors leading to enhanced hypoxia tolerance
Water quality
In our previous study (Gamperl et al.,
2001), trout of similar age/size were obtained from a rearing
facility supplied with groundwater at a constant temperature of 8°C (West
Creek Spring Trout Farm, Aldergrove, BC, Canada). In contrast, the hatchery
that provided the trout used in this study is supplied with a limited amount
of stream water that is subject to seasonal variations in temperature
(820°C) and oxygen content (minimum 5 mg O2
l1). In addition, the holding pens are arranged in series
(fry, juveniles, sub-adults, then adults), and water quality deteriorates as
maturing fish are moved away from the farm's water source. It is probable,
therefore, that selection occurred at this facility, with only hearty and/or
hypoxia-tolerant individuals surviving to adulthood. This selective pressure
would have been greatest during the summer when stream temperatures were
highest (
20°C), because high water temperatures lead to (1) increased
resting metabolic rates (Q10=1.52.0;
Brett, 1971
;
Dickson and Kramer, 1971
) and
(2) a reduction in the oxygen carrying capacity of water (1.5% for every
°C increase in temperature).
Branchial copepod infestations
The trout used in these experiments were active and well fed. However,
branchial copepod infestations (probably Ergasilis sp.) were
widespread in this population. It is possible that these parasitic
infestations reduced the amount of oxygen available to the myocardium and
other tissues, and thus magnified the influence of environmental hypoxia
(water quality) on fish survival and/or cardiac physiology. Specifically, gill
damage caused by these parasites would reduce the gill surface area available
for gas exchange, and decrease oxygen uptake from the water. It is also
possible that bleeding associated with these infestations led to a reduced
haematocrit. A low haematocrit would impair oxygen transport within the blood,
and require the heart to work harder in an attempt to maintain oxygen delivery
to the tissues during periods of aquatic hypoxia. Such an increase in
myocardial energy expenditure might have further exacerbated the effects of
reduced blood oxygen content on the cardiac muscle and promoted greater
hypoxia tolerance.
Potential adaptations mediating hypoxia tolerance
The average cardiac LDH activity measured in this study (136.0±15.2
units g1 wet mass (Fig.
7) is only slightly higher than that reported for other rainbow
trout populations (110 units g1 wet mass) if LDH activity is
adjusted to a common temperature (10°C, assuming Q10=2;
Driedzic and Gesser, 1994).
Although measurements of maximal enzyme activity may not reflect
`physiological flux' through a pathway or even a single step, our measurements
of cardiac LDH activity suggest that the enhanced myocardial hypoxia tolerance
of these in situ hearts is not due to an improved ability to generate
ATP through anaerobic glycolysis.
Although cardiac myoglobin was not measured in this study, a greater
concentration of myoglobin would increase the efficiency of oxygen use at low
PO2 levels
(Bailey et al., 1990).
Ultimately, myoglobin would help to fuel aerobic metabolism, improve energy
availability, and minimize the negative effects associated with the
accumulation of anaerobic by-products. There is evidence linking myocardial
myoglobin levels and hypoxia-tolerance in both the fish and mammalian
literature. Inter-specific differences in fish myoglobin concentration play an
important role in the maintenance of cardiac function during hypoxia
(Driedzic et al., 1982
;
Legate et al., 1998
), and rats
experience a 15% increase in cardiac myoglobin concentration following 2 to 10
weeks of hypoxic adaptation (PO =7390 mmHg;
Anthony et al., 1959
). However,
myoglobin levels were not altered following 46 weeks of hypoxic
exposure (water PO =3035 mmHg) in the eelpout
(Zoarces viviparous; Driedzic et
al., 1985
). This latter study suggests that cardiac myoglobin
concentrations do not increase in the fish heart following chronic hypoxic
exposure, and that increased levels of myoglobin in the myocardium were not
responsible for the enhanced hypoxia tolerance of the rainbow trout hearts
used in this study.
A number of sources suggest that an enhanced ability to use exogenous
glucose improves post-hypoxic recovery of maximum myocardial function. First,
hypoxia stimulates myocytes to take up glucose by recruiting glucose
transporters to the cell membrane and/or improving the function of existing
glucose transporters (Cartee et al.,
1991; Rodnick et al.,
1997
). Second, exogenous glucose enhances cardiac function during
oxygen deprivation in mammals (Apstein et
al., 1983
; Runnman et al.,
1990
), in hypoxia-tolerant eels
(Driedzic et al., 1978
;
Bailey et al., 2000
) and in the
hypoxia-adapted eelpout (Zoarces viviparous;
Driedzic et al., 1985
). The
functional protection afforded by exogenous glucose may be mediated by an
increased ability to fuel anaerobic glycolysis, as observed by Gamperl et al.
(2001
). The advantages of
elevating anaerobic glycolysis may include (1) an increased production of ATP
and (2) the maintenance of vital membrane functions (i.e. ionic balance,
membrane potential), particularly if the specific enzymes required for glucose
breakdown exist in close proximity to the cell membrane
(Runnman et al., 1990
). In
addition, exogenous glucose may further protect the in situ heart by
acting as a free radical scavenger during reoxygenation
(Hess et al., 1983
). Although
glucose uptake was not measured, it is possible that the in situ
hearts used in this study had an enhanced rate of glucose uptake from the
hypoxic perfusate compared to our previous studies
(Arthur et al., 1992
;
Gamperl et al., 2001
), and
that this enhanced the post-hypoxic recovery of myocardial function.
Alternatively, fish used in the current study might have a higher
concentration of endogenous glucose (glycogen) that promotes glycolytic
activity and post-hypoxic recovery.
Biochemical indices of myocardial damage
The release of cardiac myoglobin and changes in ventricular LDH activity
were measured to determine if these biochemical markers could be used as
direct indices of in situ cardiac cell death. These biochemical
markers were selected because both myoglobin
(Janier et al., 1994;
Kawabata et al., 1998
;
Stokke et al., 1998
) and LDH
(McKean and Mendenhall, 1996
;
Diederichs, 1997
) are released
from the mammalian heart in response to acute cardiac injury. In addition, the
release of cardiac metabolic enzymes such as LDH and creatine kinase has been
successfully measured in (1) coronary venous blood
(McKean and Landon, 1982
;
McKean and Mendenhall, 1996
),
(2) small volumes of perfusate holding in vitro heart preparations
(Ghosh et al., 2000
) and (3)
the systemic circulation (Diederichs,
1997
).
Although protein was released from the hearts following severe hypoxia,
myoglobin was not detected in the perfusate leaving these in situ
hearts, and ventricular LDH activity did not vary with the duration of severe
hypoxia (Fig. 7). It is
possible that these direct biochemical indicators of myocardial damage were
not detected because either (1) dilution of myoglobin within the large volumes
of perfusate pumped by in situ heart resulted in concentrations that
were below the detection limit of the assay, or (2) 30 min of oxygenated
reperfusion was insufficient to remove enough LDH from irreversibly damaged
myocytes so that a significant reduction in tissue concentration could be
detected. However, it is unlikely that the inadequate time for washout
explains the lack of necrosis suggested by our results for perfusate myoglobin
and myocardial LDH. McKean and Mendenhall
(1996) showed that large
amounts of LDH are released from the mammalian heart during the first 30 min
of reperfusion. Overgaard et al.
(2004
) showed that the
energetic status (total adenylates, adenylate charge, lactate, glycogen and
PCr/Cr2) of trout hearts exposed to 20 min of severe hypoxia was
not significantly different from control hearts after 30 min of oxygenated
reperfusion. Finally, anoxic and normoxic non-working myocardial pieces
(46 mg) from rainbow trout showed no difference in MTT staining until
at least 4 h of incubation (J. Overgaard and J. A. W. Stecyk, unpublished).
Thus, we hypothesize that in situ trout hearts are not irreversibly
damaged when exposed to brief (<30 min) periods of severe hypoxia at
10°C, and that myocardial dysfunction during recovery is solely a result
of `stunning'.
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Acknowledgments |
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Footnotes |
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References |
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