Effects of temperature and anoxia upon the performance of in situ perfused trout hearts
1 Department of Zoophysiology, Institute of Biological Sciences, University
of Aarhus, Denmark
2 Department of Biological Sciences, Simon Fraser University, British
Columbia, Canada
* Author for correspondence (e-mail: johannes.overgaard{at}biology.au.dk)
Accepted 13 November 2003
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Summary |
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At high oxygenation, maximum cardiac output and power output increased with temperature (Q10 values of 1.8 and 2.1, respectively) as a result of increased heart rate. Hypoxia tolerance was inversely related to temperature. At 5°C, the hearts maintained routine cardiac output throughout the 20 min period of anoxia, and maximal cardiac performance was fully restored following reoxygenation. By contrast, cardiac function failed sooner during anoxia as temperature was increased and maximal performance after reoxygenation was reduced by 25%, 35% and 55% at 10°C, 15°C and 18°C, respectively. Increased functional impairment following anoxic exposure at elevated temperature occurred even though both cardiac glycolytic enzyme activity and the rate of lactate production were increased proportionally with cardiac work. Nonetheless, there was no indication of myocardial necrosis, as biochemical and energetic parameters were generally unaffected by anoxia.
Key words: fish, trout, Oncorhynchus mykiss, temperature, cardiovascular, recovery, in situ perfusion, hypoxia, anoxia, glycolytic metabolism
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Introduction |
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Rainbow trout (Oncorhynchus mykiss) is not particularly hypoxia
tolerant, but may encounter substantial variations in oxygen availability in
both natural settings and when kept in aquaculture. When exposed to hypoxia,
rainbow trout exhibit the typical piscine cardio-respiratory response (e.g.
Randall, 1982) and, in common
with other animals, reduce their preferred body temperature during hypoxia by
seeking lower environmental temperatures
(Schurmann et al., 1991
;
Steiner and Branco, 2002
).
Hypoxia-induced hypothermia diminishes metabolism, as well as the critical
oxygen level (Schurmann and Steffensen,
1997
; Ott et al.,
1980
), and serves a protective role by alleviating the demands on
the cardiovascular system. The importance of suppressing cardiac demands is
well illustrated by the finding that the 70% reduction in force production by
anoxic myocardial strips from rainbow trout occurs faster, albeit ultimately
to the same extent, at 20°C compared with 10°C
(Hartmund and Gesser, 1992
).
While reduced temperature confers benefits to prolong the activity of anoxic
cardiac tissue by reducing metabolism and the rate of the ensuing lactic
acidosis, decreased temperature also diminishes glycolytic capacity, which may
affect the balance between energy production and energy consumption.
Using in situ perfused hearts from trout, we investigate the effects of temperature on routine and maximal cardiac performance during normoxia, anoxia and following recovery from anoxia. We tested the hypothesis that the rainbow trout heart benefits from lowered temperatures during anoxia (severe hypoxia) despite an anticipated depression of glycolytic capacity. Measurements were taken before, during and after a 20 min exposure to anoxia in preparations that were acutely exposed to 5, 15 and 18°C, as well as in preparations measured at the acclimation temperature of the fish (10°C). To mimic in vivo conditions, the hearts were set to work at increasingly higher loads with increasing temperature, and maximal cardiac performance was measured both before and following recovery from anoxia. These functional variables are related to glycolytic capacity and energetic status of the cardiac muscle, which were determined at the end of the experiment.
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Materials and methods |
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Surgical procedures
Fish were anaesthetised in an oxygenated buffered solution of tricaine
methane sulfonate (MS-222; 0.15 g l-1 MS-222 plus 0.15 g
l-1 sodium bicarbonate) and transferred to an operating table where
their gills were irrigated with oxygenated water containing diluted
anaesthetic (0.075 g l-1 MS-222 plus 0.075 g l-1 sodium
bicarbonate) at 8-10°C. Fish were then injected with 1.0 ml
kg-1 of heparinised (100 U ml-1) saline via the
caudal blood vessels, and the perfused heart preparation was prepared as
described by Farrell et al.
(1986). Briefly, an input
cannula was introduced into the sinus venosus through a hepatic vein, and
perfusion with heparinised (10 IU ml-1) saline containing 7.5 nmol
l-1 adrenaline was immediately commenced (the adrenaline
concentration was higher for 18°C hearts; see below). An output cannula
was inserted into the ventral aorta at a point confluent with the bulbus
arteriosus and tied firmly in place with a single silk thread. Finally, both
ducts of Cuvier were occluded with sutures, thereby crushing the cardiac
branches of the vagus nerve and eliminating all venous return to the heart
except for the perfusate delivered to the heart via the input
cannula. Thus, the in situ preparation isolated the heart in terms of
perfusate delivery and collection, as well as autonomic nervous control, while
leaving the pericardium intact.
Once surgery was completed (15-20 min after netting), the fish was
immersed in a temperature-controlled saline bath regulated at the test
temperature (either 5, 10, 15 or 18°C). The input cannula was attached to
an adjustable constant-pressure reservoir, and the output cannula was
connected to a constant pressure head. For routine normoxic conditions, output
pressure (Pout) was set to 4.9 kPa to simulate in
vivo ventral aortic blood pressure at rest, and input pressure to the
heart (Pin) was adjusted to give a routine cardiac output
that was physiological for the particular test temperature (approximately 12,
16, 20 and 24 ml min-1 kg-1 for 5, 10, 15 and 18°C,
respectively; Table 1)
(Kiceniuk and Jones, 1977
;
Thorarensen et al., 1996
). As
a result, Q10 of routine cardiac output and power output varied
between 1.6 and 1.8 for the different temperature intervals (average 1.7),
which resembles in vivo Q10 values (
2;
Driedzic and Gesser, 1994
). In
the absence of autonomic input, heart rate was set by the intrinsic pacemaker
and the stimulation provided by the tonic level of adrenaline in the
perfusate. The 7.5 nmol l-1 of adrenaline used for 5, 10 and
15°C hearts is similar to that of plasma in resting rainbow trout
(Milligan et al., 1989
).
However, as in previous studies of perfused rainbow trout hearts studied at
temperatures approaching their upper lethal limit
(Farrell et al., 1996
), our
preparations needed considerably higher adrenergic stimulus to maintain stable
cardiac output and heart rate at 18°C. Therefore, at 18°C the
perfusate contained 50-100 nmol l-1 adrenaline. Cardiac output was
maintained at a routine level for 20 min to allow heart function to stabilise
and to ensure temperature equilibration with the saline bath. Hearts that
required a Pin of >0.05 kPa to reach routine cardiac
flow rate (
) were discarded because
this indicates either a dysfunctional pericardium or improper input cannula
placement in the preparation. However, two out of seven hearts at 18°C
were allowed to start with marginally higher input pressures because these
preparations appeared normal in all other respects. Adjustments of
Pin were made continuously to maintain routine
.
|
The perfusate contained 124 mmol l-1 NaCl, 3.1 mmol
l-1 KCl, 0.93 mmol l-1 MgSO4.7H2O,
2.52 mmol l-1 CaCl2.2H2O, 5.6 mmol
l-1 glucose, 6.4 mmol l-1 TES salt and 3.6 mmol
l-1 TES acid. The TES buffer simulates the buffering capacity of
trout plasma and the normal change in blood pH with temperature
(pK/dT of 0.016 pH units deg.-1). The perfusate was
renewed every 20 min, as adrenaline rapidly degrades. The coronary circulation
that normally supplies the outer 30-40% of the rainbow trout heart (Farrell et
al., 1986
,
1988a
) was not perfused in our
preparation. To compensate, the perfusate was gassed with 100% O2,
and Gamperl et al. (2001
)
argued that this level of oxygenation can supply a sufficient amount of
O2 to the outer myocardial layer because the O2 tension
in the perfusate is more than 10x that routinely found in venous blood
in rainbow trout (Farrell and Clutterham,
2003
). As a result, maximal performance of the in situ
perfused rainbow trout heart is comparable with, or even higher than, the
maximal performance measured in vivo (Farrell et al.,
1986
,
1991
). For the anoxic (severe
hypoxic) exposures, the perfusate reservoir was gassed with 100% N2
for at least 1 h, which resulted in a partial oxygen pressure
(PO2) of <0.5 kPa. Oxygen transfer between
the surrounding bath and the heart was minimised by covering the preparation
with a loose-fitting plastic lid and bubbling the saline bath with 100%
N2, beginning 5 min before the onset of anoxia.
Experimental protocols
The experimental protocols were designed to examine the effects of
temperature on normoxic, anoxic and post-anoxic performance. Maximal and
routine performance was assessed, and recovery of cardiac performance was
measured after exposure to a standardised 20-min period of anoxia. During
anoxia, Pin was gradually raised to, but not above, 0.25
kPa in an attempt to maintain routine
. If 0.25 kPa was insufficient, flow
was allowed to decline. Following anoxia, Pout was reduced
to 1.0 kPa for 10 min to aid the recovery of cardiac output.
Fish were randomly assigned to one of five experimental treatment groups (N=6 or 7). The fish tested at 5°C were significantly larger than those tested at 18°C, but the ratio between ventricle mass and body mass (0.089±0.002%) was similar for all temperature groups (Table 1). The five experimental treatments were as follows: (A) Oxygenated control tested at 10°C; these hearts were only exposed to oxygenated saline and were intended to reveal any decay in cardiac performance over time; (B-E) anoxic exposure; hearts were tested at 5, 10, 15 or 18°C, using routine cardiac outputs of 12, 16, 20 or 24 ml min-1 kg-1, respectively. The various protocols, which all lasted 140 min, are depicted schematically in Fig. 1. After stabilisation for 20 min at the routine cardiac output, maximum cardiac output was assessed under normoxic conditions (see maxtest protocol below). Subsequently, the hearts were allowed to recover at routine cardiac output for 55 min, whereupon they were exposed to 20 min of anoxia, followed by 30 min of normoxic recovery before a second maxtest was performed.
|
Assessment of cardiac performance and viability
Assessment of cardiac performance was based on repeated measurements of
routine and maximum cardiac performance, while viability was based on
biochemical measures. Maximal cardiac performance (maxtest) was determined by
increasing Pin up to 0.45 kPa to maximise stroke volume
and reach maximal flow rate
(max). Then,
Pout was progressively increased to attain maximal power
output (POmax). After the maxtest,
and Pout were
returned to routine levels. Routine cardiac performance was measured
continuously throughout the experiment and maximal performance was assessed at
the start of the experiment during normoxia before other experimental
manipulations (maxtest1) and repeated 30 min after anoxia
(maxtest2) (Fig. 1).
In this way, each heart served as its own control for statistical
analysis.
Perfusate from the outflow was sampled at 2 min intervals during the 20 min
anoxic challenge (arrows in Fig.
1) tomeasure lactate efflux from the heart. Samples were stored at
-80°C until analysis with an automated YSI 2300 Stat Plus glucose and
L-lactate analyser (YSI Inc., Yellow Springs, OH, USA). Lactate
efflux rate (nmol lactate g-1 ventricle min-1) was
calculated as:
![]() | (1) |
Maximal rate of lactate efflux during anoxia was defined as the maximal
value measured at stable heart rate (fH), and lactate
measurements were discarded if fH or
were unstable or approached zero.
Thus, due to rapidly failing flow during anoxia at 18°C, no lactate efflux
rates are reported at that temperature.
Myocardial enzyme activities and high-energy phosphate concentrations were
measured to evaluate myocardial necrosis and to determine whether energy
balance had been restored after the various treatments. Ventricular tissue was
sampled 5 min after maxtest2, i.e. approximately 40 min after the
conclusion of the anoxic period. Tissue was quickly excised and freeze-clamped
with aluminium tongs precooled in liquid nitrogen, split into several pieces
that were weighed and stored at -80°C. Biochemical analysis was performed
on 50-200 mg muscle samples homogenised in 50 volumes of 50% glycerol in 20
mmol l-1 sodium phosphate buffer (pH 7.4), 5 mmol l-1
ß-mercaptoethanol, 0.5 mmol l-1 EDTA and 0.02% bovine serum
albumin (BSA). Activities of the glycolytic enzymes lactate dehydrogenase and
pyruvate kinase were measured at 20°C as described by Chi et al.
(1983) and Christensen et al.
(1994
), respectively, with some
modifications. The lactate dehydrogenase reaction solution contained 100 mmol
l-1 imidazole adjusted to pH 7.4, 1 mmol l-1 sodium
pyruvate, 0.3 mmol l-1 NADH and 0.05% BSA. Pyruvate kinase activity
was measured in a reaction solution consisting of 50 mmol l-1
triethanolamine, 75 mmol l-1 KCl, 8 mmol l-1
MgSO4, 0.8 mmol l-1 phosphoenolpyruvate, 1 mmol
l-1 ADP, 0.3 mmol l-1 NADH and 60 U ml-1
lactate dehydrogenase that was adjusted to pH 7.5. Both enzymatic activities
were expressed relative to ventricular protein content, which was measured
from similar homogenates (without BSA) using Sigma kit no. 690-A (Sigma
Chemicals, Oakville, ON, Canada). In addition to the measurements of enzymatic
activity at 20°C, enzymatic activities were also measured at 5, 10 and
15°C in six randomly chosen samples to obtain the temperature sensitivity
of these glycolytic enzymes.
Myocardial high-energy phosphates were measured using high-preformance
liquid chromatography (HPLC; Bøtker
et al., 1994). Briefly, a 30-60 mg piece of ventricle was
homogenised in 1.6 ml of 0.42 mol l-1 perchloric acid (PCA) in a
glass-glass homogeniser. The homogenate was then centrifuged for 10 min at
3400 g, and the supernatant was separated into two 200 µl
portions. One portion was used for measurement of creatine compounds
(phosphocreatine and creatine) and was neutralised with 100 µl KOH (1 mol
l-1), while the other, used for subsequent measurement of
adenylates (ATP, ADP and AMP), was neutralised with 100 µl of
KHCO3 (2 mol l-1) and Tris (0.1 mol l-1).
After neutralisation, both portions were kept on ice for 10 min to ensure
precipitation of perchlorate. The neutralised portions were then centrifuged
for 5 min (3400 g) and the supernatant stored at -80°C
until further analysis. Creatine compounds and adenylate compounds were
separated using HPLC (Waters, Milford, MA, USA) with a 10 cm crompack C18
microsphere-column of 3 µm particle size (Varian, Palo Alto, CA, USA).
Creatine compounds were measured at a wavelength of 210 nm using a mobile
phase of aqueous buffer containing 0.02 mol l-1
KH2PO4 and 2.3 mmol l-1 tetrabutyl ammonium
hydrogen sulphate (TBAHS) run at 1.5 ml min-1. Adenosine
nucleotides were measured at a wavelength of 254 nm using a mobile phase of
25% methanol and 75% aqueous buffer containing 0.06 mol l-1
KH2PO4 and 0.011 mol l-1 TBAHS run at 1 ml
min-1. Cardiac energy status was expressed as adenylate energy
charge: ([ATP]+ 0.5[ADP])/([ATP]+[ADP]+[AMP]) and phosphorylation potential:
[PCr]/[Cr]2 (Meyer,
1988
).
Muscle lactate was measured spectrophotometrically as described by Lowry
and Passonneau (1972) from the
same homogenates as those used for adenylate measurements. Briefly, lactate
was measured from the change in absorbance following addition of lactate
dehydrogenase (15 U ml-1) in a reaction buffer containing 50 mmol
l-1 glutamic acid, 50 mmol l-1
2-amino-2metyl-propanolol, 1.5 mmol l-1 NAD and glutamate-pyruvate
transaminase (3 U ml-1). Muscle glycogen was measured according to
Bergmeyer (1983
). After
homogenisation in 0.6 mol l-1 PCA, glycogen in the homogenate was
transformed to glucose by incubation for 3 h at 35°C in an acetate buffer
(0.1 mol l-1, pH 4.8) containing amyloglucosidase. Subsequently,
glucose was measured spectrophotometrically in a 0.3 mol l-1
triethanolamine buffer containing 4 mmol l-1 MgSO4, 0.4
mmol l-1 ATP and 0.4 mmol l-1 NADP before and after
addition of hexokinase and glucose-6-phosphate dehydrogenase. Muscle glycogen
content is, therefore, presented as a glucose concentration and calculated as:
[glucose] in homogenate - [glucose] in homogenate supernatant (no glycogen
transformation).
Instrumentation set-up and data analysis
An in-line electromagnetic flow probe (Zepeda Instruments, Seattle, WA,
USA) was used to record ventral aortic flow
(), and disposable pressure
transducers (model DPT-6100; PVB Medizintechnik, Hauptstraße, Germany)
were used to measure inflow and outflow pressures. Pressure transducers were
calibrated daily against a static water column and referenced to the surface
of the saline bath regularly. Pressure transducer signals were amplified with
a Senselab amplifier (Somedic Sales AB, Hörby, Sweden), while the flow
signal was amplified using a DC amplifier (Gould, Cleveland, OH, USA). All
signals were recorded at 5 Hz on an in-house programmed computer-assisted data
acquisition system (National Instruments, Austin, TX, USA), and block averages
were calculated every 5 s. The measured values for Pin and
Pout were adjusted for cannulae resistances so that the
reported pressures represented those in the sinus venosus and in the bulbus
arteriosus, respectively. fH was measured by counting
pressure pulses over 10 s periods. Ventricular power output (PO; mW
g-1 ventricle) was calculated as:
![]() | (2) |
Stroke volume (VS) was calculated as:
![]() | (3) |
Statistics
All statistical analyses were performed using SigmaStat for Windows 2.03
(SPSS Inc., Chicago, IL, USA). One-way analyses of variance (ANOVAs) were used
to compare parameters between treatment groups, including: (1) body and
ventricular mass; (2) Pin required to attain routine
cardiac output; (3) routine values of VS and
fH; (4) maximal cardiac performance during
maxtest1; (5) relative recovery of function during
maxtest2; (6) biochemical status of the muscle samples obtained at
the end of the experiments (including pyruvate and lactate dehydrogenase
activity at 20°C, protein, adenylate, creatine, glycogen and lactate
content, adenylate energy charge and [PCr]/[Cr]2). A nonparametric
test (Mann-Whitney) was used when data were not normally distributed.
Repeated-measures ANOVAs were performed for comparisons of: (1) maximum
cardiac performance within a treatment group at the initial and second test
and (2) Pin, PO, ,
VS, and fH during the 20 min anoxic
challenge relative to the control (pre-anoxic) conditions. Unless otherwise
stated, statistical differences were identified using a Bonferroni
post-hoc test, and P<0.05 was used as the level of
statistical significance. All data are reported as means ±
S.E.M.
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Results |
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Input pressure was increased gradually during the experiment to maintain routine cardiac output (Fig. 2). After anoxia, it was often necessary to increase Pin, and the increase in Pin was particularly pronounced at 18°C (Fig. 2A).
|
Cardiac performance during anoxia
Normoxic cardiac performance increased progressively with elevated
temperature, while cardiac anoxia tolerance was inversely related to
temperature (Figs 2,
3). Hearts maintained cardiac
output throughout anoxia at 5°C (Fig.
2E), but anoxia tolerance decreased with increased temperature, as
indicated by the pronounced and rapid decline of cardiac output in spite of
the increased input pressure (Fig.
2). Routine cardiac output was significantly reduced after 15 min
and 20 min of anoxia at 15°C and 10°C, respectively, and significantly
depressed after only 5 min of anoxia at 18°C. Nevertheless, cardiac output
was similar at the end of anoxia (approximately 5 ml min-1
kg-1) in all groups at 10°C or warmer.
|
Cardiac performance after anoxic recovery
All preparations were allowed 30 min to recover from anoxia before
maxtest2. Maximal performance of the oxygenated control at 10°C
was unchanged over time (Fig.
3). While maximal performance was not affected by anoxia at
5°C, the ability of the in situ perfused heart to recover from
anoxic exposure was inversely related to temperature and the reduction of
maximal performance was 55% at 18°C
(Fig. 3). The reduction in
maximal cardiac performance following recovery from anoxia that was observed
at temperatures above 5°C was mainly caused by a reduction in stroke
volume, while post-anoxic heart rate was only marginally affected
(Fig. 3C,D). Curiously, the
absolute levels for maximum cardiac output and power output of the recovered
hearts did not vary significantly with temperature and were 40 ml
min-1 kg-1 and
3.8 mW g-1 ventricle,
respectively, at all temperatures (Fig.
3A,B). Heart rate was unaffected except at 15°C
(Fig. 3D).
The effect of acute temperature changes on glycolytic metabolism
When measured at 20°C, maximal enzymatic activities of lactate
dehydrogenase (LDH) and pyruvate kinase (PK) were similar in all experimental
groups (Table 2), indicating
that all hearts possessed similar capacities for glycolytic energy production.
However, maximal enzymatic activity was temperature dependent
(Fig. 4) with Q10
values of 1.9 and 2.3 for LDH and PK, respectively, between 5°C and
20°C. The Q10 values were considerably higher for both enzymes
between 5°C and 10°C compared with between 10°C and 20°C
(Q10=2.5 vs 1.6 for LDH and 5.0 vs 1.6 for
PK).
|
|
The temperature dependency of the glycolytic capacity was reflected in a temperature dependency of lactate efflux from the anoxic perfused hearts (Fig. 5). We estimate the highest rates of lactate production to be 2.0, 2.7 and 4.1 µmol min-1 g-1 ventricle at 5, 10 and 15°C, respectively (Fig. 5), which yielded a Q10 of 2.1, which is comparable with the Q10 of the glycolytic enzymes. (We were unable to quantify lactate production reliably at 18°C as cardiac output failed to stabilize almost immediately after the onset of anoxia.)
|
Biochemical and energetic state of the myocardium following recovery
There were only very limited differences in myocardial state after recovery
from anoxia. Enzymatic capacities of PK and LDH, glycogen content and total
creatine were similar between temperature groups
(Table 2), indicating a lack of
cellular rupture and absence of energetic depletion of the myocardium. In
general, energetic status was similar between groups, although adenylate
charge was significantly lower at 18°C compared with 5°C, coupled with
significantly lower total adenylates at 18°C. Hearts at 18°C were
characterised by a higher level of muscle lactate compared with those at
5°C and 15°C, which may indicate that myocardial homeostasis had not
fully recovered at 18°C.
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Discussion |
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Maximal flow and power output reported in our study are generally similar
to those in previous in situ and in vivo studies. While our
values for maximal cardiac performance at 10°C and 15°C are slightly
lower than those in previous studies
(Kiceniuk and Jones, 1977;
Farrell et al., 1991
,
1996
), performance at 18°C
closely resembles that reported by Farrell et al.
(1996
). Conversely, our values
at 5°C are considerably higher than those reported at 4°C in
vivo by Taylor et al.
(1996
). However, direct
comparison is difficult because the level of adrenergic stimulation differs
among studies. Here, we used a tonic level of adrenaline (7.5 nmol
l-1) at 5, 10 and 15°C to simulate resting in vivo
plasma concentrations (Milligan et al.,
1989
). The importance of tonic adrenaline and its variable
contribution with temperature was illustrated at 18°C, where adrenaline
concentration had to be increased to 50-100 nmol l-1 to avoid
cardiac arrhythmias. Similar problems have been encountered previously at
temperatures above 18°C with rainbow trout hearts
(Farrell et al., 1996
).
Here, heart rate increased with a Q10 of 1.7 from 5°C to
18°C, as in vivo (Taylor et
al., 1996). In acclimated rainbow trout, the positive chronotropic
effect of temperature is reduced at high temperatures, and Farrell et al.
(1996
) reported a
Q10 of 1.3 for heart rate between 15°C and 22°C for in
situ perfused hearts. We observed a much higher Q10 (1.9) for
heart rate between 15°C and 18°C, but this may have been influenced by
the 10-fold higher adrenaline concentration applied to the hearts at 18°C
as well as the result of an acute temperature change versus
temperature acclimation. Recently, using a similar perfused heart set-up,
Blank et al. (2002
) found that
the Q10 for heart rate in yellowfin tuna (Thunnus
albacares) was considerably higher than in rainbow trout (approximately 3
at 10-25°C), but stroke volume was halved with each 10°C increase in
temperature. The reduced routine stroke volume in tuna may reflect either a
negative inotropic effect or a reduced cardiac filling time during diastole at
the very high heart rates of this species.
Maximum stroke volume measured during the initial maxtest was not affected
by temperature (Table 1) and
this contrasts with results for acclimated rainbow trout, where maximum stroke
volume was reduced at acclimation temperatures above 15°C
(Farrell et al., 1996). It is
possible that the higher adrenergic stimulation at 18°C explains part of
this difference, which would be further exacerbated by the low sensitivity to
catecholamines in rainbow trout acclimated to high temperature
(Keen et al., 1993
). The
unchanged maximum stroke volume is, however, in accord with the unchanged
twitch force of trout ventricular strips following acute temperature changes
(Hartmund and Gesser, 1992
;
Hove-Madsen, 1992
), although
negative inotropy with increased temperature was observed at low temperatures
in rainbow trout (Hove-Madsen,
1992
).
Cardiac performance during anoxia
The major finding of the present study was that anoxia tolerance and
recovery of the in situ perfused hearts performing at physiologically
realistic workloads were inversely related to temperature. Hearts studied at
5°C maintained routine cardiac output throughout 20 min of anoxia and
fully recovered maximal performance upon reoxygenation. By contrast, hearts
failed within the first 10 min of anoxia and suffered a 55% loss of maximal
performance after recovery at 18°C. The loss of function during anoxia at
elevated temperature occurred in spite of an increased lactate efflux and
glycolytic enzyme activities with elevated temperature. Despite the
significant reduction in maximal cardiac performance above 5°C, there was
no indication of myocardial necrosis, as biochemical and energetic parameters
were generally unaffected, although some minor changes may have occurred at
18°C (Table 2). These
results support our hypothesis that low temperature provides benefits towards
myocardial hypoxia tolerance. Here, we show that, when devoid of a myocardial
oxygen supply, routine cardiac performance can be maintained for a brief
period at 5°C and shortly afterwards perform at a maximum level when
oxygenation is restored. These results indicate that rainbow trout may have a
physiological advantage in exhibiting behavioural hypoxic hypothermia when
exposed to hypoxia (Schurmann et al.,
1991).
The inability of the rainbow trout heart to maintain performance during
anoxia is most likely caused by insufficient anaerobic energy production.
Consistent with our study, Arthur et al.
(1992) reported that glycolytic
capacity of perfused trout hearts at 16°C could supply only half of the
energetic requirement of oxygenated hearts working at routine conditions.
Efficiency of rainbow trout hearts is unaffected by severe hypoxia
(Arthur et al., 1992
; J.
Overgaard and H. Gesser, manuscript submitted). Consequently, anoxia
substantially reduces phosphocreatine concentration in perfused hearts and
cardiac strips (Arthur et al.,
1992
; Hartmund and Gesser,
1996
), and cardiac failure appears to be caused by increased
levels of intracellular inorganic phosphates and reduced intracellular pH
(Allen et al., 1985
;
Godt and Nosek, 1989
;
Arthur et al., 1992
). This is
supported by the observation that maximum power output and intracellular pH of
perfused rainbow trout heart decrease when the extracellular perfusate is made
acidic (Farrell et al., 1986
,
1988b
). Elevated adrenaline
concentration can, however, restore performance of an acidotic heart without
intracellular pH being restored (Farrell
and Milligan, 1986
).
Maximal lactate efflux had a Q10 of 2.1 between 5°C and 15°C and, similarly, overall Q10 values of PK and LDH were 1.9 and 2.3, respectively. Glycolytic capacity, therefore, increases as least as much as routine cardiac performance (Q10=1.7). The increased sensitivity to anoxia at high temperature, therefore, cannot be ascribed to a reduction in glycolytic capacity relative to energy requirements. Instead, it seems that the increased sensitivity is caused by a higher rate of accumulation of waste products with increased temperature, so the intracellular milieu, particularly intracellular phosphate and pH, is disturbed sooner.
Post-anoxic recovery
While several studies have examined the effects of severe hypoxia and
anoxia on cardiac function in fish, few studies have previously described
post-anoxic recovery of whole working hearts (however, see
Gamperl et al., 2001; J.
Overgaard, J. A. W. Stecyk, H. Gesser, K. Gamperl, T. Wang and A. P. Farrell,
manuscript submitted). Our results show a marked effect of temperature on
post-anoxic recovery, where hearts at 18°C only recover 45% of initial
performance while hearts were unaffected by 20 min of anoxia at 5°C. The
reductions in post-anoxic performance at temperatures above 5°C can mainly
be attributed to reductions in the contractile performance of the hearts, as
it is stroke volume rather than heart rate that is reduced following recovery
from anoxia. However, the decreased contractile performance of the hearts
after anoxia at high temperature cannot be directly explained from impaired
myocardial cellular status. Thus, we found no indication of either cellular
necrosis or rupture, as both enzyme activity and concentrations of creatine
products were similar in all experimental groups
(Table 2). Similarly, with the
exception of a significant difference in adenylate charge between 18°C and
5°C, there were no significant differences in energetic state between
groups, and the values of adenylate charge and phosphorylation potential were
similar to those of normoxic trout hearts
(Arthur et al., 1992
; J.
Overgaard and H. Gesser, manuscript submitted). Moreover, the reduction in
maximal cardiac performance is unlikely to be caused by energy depletion, as
glycogen levels were recovered to a level close to that previously reported
for normoxic rainbow trout (Gesser,
2002
). The only notable differences were higher levels of muscle
lactate and lower total adenylates at 18°C. Lactate levels after anoxia at
18°C were, nevertheless, only moderately higher and unlikely to represent
a significant intracellular acidosis. A decrease in total adenylates at
18°C was also reported for anoxic myocardial strips with an inhibited
glycolysis (Hartmund and Gesser,
1996
), and it is possible that some of the adenylates are lost as
adenosine during anoxia. While increased muscle lactate levels could be due to
increased glycolytic metabolism during oxygenated perfusion, they may also
represent residual lactate from the preceding anoxia. It is possible that
lactate accumulated to higher levels at 18°C because the hearts failed
early into the anoxic period and had rather low cardiac output during the
remaining part of anoxia. The lower recovery of routine performance would also
attenuate lactate removal after anoxia.
Given the general lack of cell death and disruption of myocardial energetic
and enzymatic status in the failing hearts, we propose that decreased
post-anoxic performance is due to myocardial stunning rather than necrosis.
Myocardial stunning is defined as the mechanical dysfunction that persists
after reoxygenation (reperfusion) despite the absence of irreversible damage
(Bolli and Marban, 1999). The
severity of myocardial stunning in mammals is associated with both the
duration of flow deprivation and temperature, and the decreased contractility
is thought to stem from a reduction in Ca2+ responsiveness caused
by damage of the contractile apparatus of oxygen radicals and/or
Ca2+ overload (Bolli and Marban,
1999
). We suggest increased levels of oxygen radicals to be the
primary candidate of stunning in trout, as Ca2+ overload only
induces modest reductions in twitch force in ectothermic vertebrates compared
with mammals (Poupa et al.,
1985
).
Conclusion
In the present study, we found that maximal and routine cardiac performance
increase between 5°C and 18°C with a Q10 of 1.7-2.1 due to
increased heart rate, while maximal and routine stroke volume were unaffected
by acute temperature changes. Anaerobic capacity of the heart, as assessed
through measurements of lactate efflux and enzymatic capacity, increased at a
similar, but slightly higher, rate to that of routine cardiac output. Even so,
20 min of anoxia resulted in cardiac failure at 10, 15 and 18°C, with the
rate of development of cardiac failure and the extent of residual impairment
upon reoxygenation both being temperature dependent. By contrast, hearts at
5°C were not significantly affected by anoxia. Thus, the consequences of
anoxia are much more severe at high temperature because the hearts are working
at higher rates. We suggest that increased levels of inorganic phosphates and
protons cause cardiac failure during anoxia and that the accumulation of waste
products is exacerbated once flow can no longer be maintained. Thus, even
though glycolytic capacity decreases more than cardiac work with an acute
decrease in temperature, the rate at which myocardial homeostasis is disturbed
is slower due to the low metabolism. While energetic state was normalised
after recovery from anoxia, post-anoxic cardiac performance was indirectly
related to the test temperature. We propose that the mechanism underlying
post-anoxic failure is myocardial stunning and that the degree of myocardial
stunning is influenced by the degree of cardiac failure during the preceding
anoxic period. Given these results, we conclude that fish exposed to anoxia
have a clear physiological advantage in exhibiting behavioural hypoxic
hypothermia, as this should aid them to endure and recover from hypoxic
insults.
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References |
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