Vol. 273, Issue 6, R2046-R2054, December 1997
Effects of hypoxia and low temperature on substrate fluxes in
fish: plasma metabolite concentrations are misleading
François
Haman,
Georges
Zwingelstein, and
Jean-Michel
Weber
Biology Department, University of Ottawa, Ottawa, Ontario, Canada
K1N 6N5
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ABSTRACT |
Oxygen levels and temperature can
fluctuate rapidly in aquatic environments. Even though the effects of
environmental stresses on fish metabolism have been studied
extensively, information on fuel kinetics is extremely limited because
it relies almost exclusively on changes in substrate concentrations.
The turnover rate of nonesterified fatty acids (NEFA) has never been
measured in fish. Therefore, our goal was to quantify glucose and NEFA fluxes in rainbow trout acutely exposed to severe hypoxia (25% O2 saturation) or low temperature
(6°C for fish acclimated to 15°C) by performing continuous
infusions of
6-[3H]glucose and
1-[14C]palmitate in
vivo. Results show that hypoxia causes a 53% decrease in NEFA turnover
rate, together with a transient increase in hepatic glucose production,
whereas a rapid drop in temperature induces equivalent declines in
glucose, NEFA, and oxygen fluxes [temperature coefficient 
2]. More importantly, kinetic changes in glucose and NEFA fluxes
are not accompanied by interpretable changes in the plasma
concentrations of these metabolites. Thus using concentration changes
to draw conclusions about fluxes must be avoided.
oxidative substrate fluxes; glucose kinetics; fatty acid kinetics; oxygen availability; fish metabolism; continuous tracer infusions; rainbow trout; Oncorhynchus mykiss
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INTRODUCTION |
HYPOXIA AND TEMPERATURE stresses can frequently occur
in aquatic environments (6, 14, 30), and their effects on fish metabolism have been the subject of many investigations. Reduced oxygen
availability is known to have profound metabolic consequences, particularly for hypoxia-sensitive teleosts such as rainbow trout (2, 7, 8, 31, 36). Even though this species is able to increase oxygen
extraction when exposed to prolonged hypoxia (9, 22), aerobic pathways
cannot maintain normal ATP turnover on their own, and anaerobic
metabolism is stimulated (2, 18, 30). Acute exposure to low
environmental temperature can compromise the normal function and the
structural integrity of fish cells (16) via changes in membrane
fluidity (13), catalytic rates of enzymes (10, 17), and diffusive
processes (26, 37). Unfortunately, our present understanding of the
integrative response to these stresses at the whole animal level is
still very primitive because it is based almost entirely on
measurements of metabolite concentrations. Changes in concentration do
not provide reliable inference about metabolite fluxes because they
merely reflect transient differences between rates of appearance
(Ra) and disappearance (Rd) (33, 39).
Therefore, the same change in concentration can be elicited by
altering Ra, Rd, or both simultaneously,
and large changes in flux can also occur while
concentration stays perfectly steady. Furthermore, the effects of
hypoxia on plasma metabolite concentrations of fish are unclear, and
rapid temperature changes have not been investigated. Some authors
report that hypoxia causes an increase in plasma glucose (15, 31) and
nonesterified fatty acids (NEFA) levels (23), whereas others show no
change in glucose (8) or a decrease in NEFA (31). It is clear that the
metabolic response of teleosts to environmental stresses can only be
captured if underlying changes in the turnover rates of energy
substrates are quantified directly. Glucose flux has only been measured
by bolus injection before and after exposure of trout to hypoxia (8),
and NEFA fluxes have never been measured in fish (33). Therefore, the
goal of this study is to quantify the turnover rates of glucose and
NEFA in rainbow trout acutely exposed to hypoxia or low temperature,
using modern methods of continuous isotope infusion (12) to obtain a
time course of changes in metabolic fuel kinetics.
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METHODS |
Animals
Rainbow trout (Oncorhynchus mykiss,
Walbaum) of both sexes were purchased from Linwood Acres Trout Farm
(Campbellcroft, Ontario, Canada) and held in 550-liter flow-through
tanks. They were kept in dechloraminated well-oxygenated water under a
12:12-h light-dark photoperiod and were divided in two groups. The
first group was kept at 10°C and used for hypoxia experiments (937 ± 53 g, n = 8), whereas the second
group was kept at 15°C and used for cold exposure experiments (714 ± 44 g, n = 8). The animals were
acclimated to these conditions for at least 1 mo before experiments
were run, and they were fed Purina trout chow three times a week until satiation.
Catheterizations
Two PE-50 catheters (Intramedic, Clay-Adams) were implanted in the
dorsal aorta under
ethyl-N-aminobenzoate sulfonic acid anesthesia (MS-222) as described previously (12). After surgery, the
animals were allowed to recover overnight in opaque Plexiglas chambers
(60 × 16 × 18 cm) supplied with the same quality water as
in their respective acclimation tanks at a flow rate of 5-6 l/min.
Continuous infusions of radiolabeled metabolites were carried out in
the same chambers, and only the animals showing hematocrits higher than
20% after recovery from surgery were used in experiments.
Metabolic Rate Measurements
During hypoxia and cold exposure experiments, oxygen consumption
(
O2)
was measured by stopping external water supply for periods of 10 min
and by recycling the same water within a 15-liter closed system.
Particular care was taken to eliminate air bubbles and to avoid
exchange between recirculated water and atmospheric air. Water flow
rate through the chamber remained at 5-6 l/min during the
metabolic rate measurements. For each 10-min measurement, O2 concentration was recorded
every minute with a calibrated oxygen electrode (Oxyguard, Handy MK
III, Valox). Chamber volume was selected to provide an optimal ratio of
fish size to respirometer volume (28). The rate of
O2
was calculated with the closed system equation of Steffensen (28).
After each measurement, return to prevailing experimental
PO2 was
accomplished by stopping recirculation and flushing the chamber for 5 min with normoxic water or hypoxic water (low
PO2 period
in hypoxia experiments).
Continuous Isotope Infusions
The infusion procedure used in these experiments has been detailed
elsewhere (12). The infusate was prepared daily with 1-[14C]palmitate
(Amersham 1.85-2.2 GBq/mmol) and
6-[3H]glucose (New
England Nuclear 1.6 TBq/mmol or Amersham 1.1 TBq/mmol). Trout plasma
was collected from donor individuals of the same batch of fish and used
as a source of lipid-binding proteins. 1-[14C]palmitate
supplied commercially in ethanol was mixed with 600 µl plasma and
well agitated before addition of
6-[3H]glucose
dissolved in Cortland saline (38). After overnight recovery from
surgery and while the animal was at rest, a priming dose of
6-[3H]glucose
equivalent to 90 min of infusion was injected before the infusion of
the
1-[14C]palmitate/6-[3H]glucose
mixture with a calibrated syringe pump (Harvard Apparatus, South
Natick, MA) at 1 ml/h was started. Infusion rates ranged between
~155,000 and 410,000 disintegrations per minute
(dpm) · kg
1 · min1
for each isotope. In all experiments, baseline glucose and fatty acid
kinetics were determined by drawing 400-µl blood samples 40, 50, and
60 min after the infusion was started. Five additional samples (400 µl each) were then taken during hypoxia or cold exposure. All samples
were centrifuged immediately, and the plasma was separated and stored
at 
20°C until analysis.
Hypoxia Experiments
After infusing the radiotracers for 1 h under normoxic conditions, the
oxygen content of the chamber was gradually decreased from 100 to 25%
saturation in 20 min by bubbling nitrogen through a column containing
glass beads. Blood samples were collected when water
PO2 reached
60 Torr (38% saturation) and 40 Torr (25% saturation) as well as 30, 60, and 90 min after reaching 25% saturation. O2
was measured under normoxic conditions and 1 h after reaching 25%
saturation.
Cold Exposure Experiments
After infusing the isotopes for 1 h at 15°C, water temperature was
gradually lowered to 6°C over 20 min. Blood samples were collected
when the temperature reached 10 and 6°C, as well as 30, 60, and 90 min after reaching 6°C.
O2
was measured three times under baseline conditions (15°C) and three
times during the 6°C exposure period.
Plasma Sample Analyses
Glucose and lactate concentrations were measured at 340 nm on a Beckman
DU 640 spectrophotometer (1). Total NEFA concentration was determined
with an analytic test-kit (NEFA C, Wako Chemicals, Osaka, Japan).
Measurement of standards for all the NEFA present in trout plasma
revealed that this test kit underestimates the concentration of 22:6 by
25%, and total NEFA concentration was corrected accordingly. The
fractional distribution of individual fatty acids to total NEFA
concentration was measured on a Hewlett-Packard 5890 series II gas
chromatograph with HP 7673 autosampler and flame ionization detector
after extraction and methylation as described previously (19, 29).
Following plasma extraction with hexane:isopropanol (3:2 vol/vol) and
centrifugation, 1 ml supernatant was counted to measure total
14C activity and partial
[3H]glucose activity.
Tritium activity remaining in the pellet was measured by dissolving it
in 1 ml NaOH (1 N) and 10 ml water. Each vial was then acidified with
glacial acetic acid (17.5 M) before counting to avoid
chemiluminescence. Palmitate activity was obtained by multiplying total
14C activity found in the
hexane:isopropanol mixture by percent activity found in the NEFA
fraction after separation of the different plasma lipids by thin-layer
chromatography (TLC). Briefly, plasma lipids dissolved in
benzene:methanol (2:1 vol/vol) were applied on silica gel plates (60 F254, Merck, Germany) and developed with heptane:isopropyl ether:acetic
acid (60:40:4). Individual lipid fractions (phospholipids + monoacylglycerol, diacylglycerol + cholesterol, NEFA, triacylglycerol,
esterified fatty acids, and cholesterol esters) were then scraped
separately in scintillation vials containing 4 ml ethanol:water (1:1)
and counted. All scintillation counting was performed in ACS-II
(Amersham) on a Tri-Carb 2500 counter (Packard, Canada).
Calculations and Statistics
Turnover rates were calculated with the equations of Steele (27),
assuming a volume of distribution of 50 ml/kg for glucose when the
non-steady state equation was used. Throughout this article, we have
used the term "flux rate" as a synonym for turnover rate, Ra, or
Rd. The turnover rate of total
NEFA was calculated by dividing palmitate turnover rate by the
fractional contribution of palmitate to total NEFA concentration.
Changes in metabolite concentrations, specific activities, and flux
rates over time were assessed by two-way analysis of variance (ANOVA),
and differences between Ra and
Rd values at individual sampling
times were evaluated by the Bonferroni
t-test. In the cold exposure
experiments, temperature coefficient
(Q10) values were calculated
with the equation of Van't Hoff using average 15°C measurements
(baseline) and the last values measured at low temperature. All values
presented are means ± SE (n = 8)
unless indicated otherwise.
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RESULTS |
Hypoxia Experiments
Water PO2, metabolic
rate, and plasma lactate.
Figure 1 shows changes in water
PO2 (line)
and metabolic rate (histogram) before and after 90 min of exposure to
hypoxia at 25% oxygen saturation. Water
PO2
decreased from 153 Torr (96% saturation) to 39 Torr (25% saturation)
in 20 min, but this change did not cause a significant decline in
metabolic rate (P = 0.29). O2
averaged 41.8 ± 6.1 µmol · kg1 · min1
throughout the experiments.
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Fig. 1.
Partial pressure of oxygen
(PO2) in
water during hypoxia experiments given as mean values (thick line) ± SE (thin lines) (n = 8). Also
shown as histogram are mean rates of oxygen consumption
(O2) ± SE in rainbow trout under normoxic and hypoxic conditions
(n = 8).
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Observed activity levels were minimal during the measurements, except
for a few seconds when water
PO2 reached
~80 Torr. After this brief episode, each animal stopped moving and
started hyperventilating until the end of the experiment. Mean plasma
lactate concentration was 1.1 ± 0.11 mM under normoxic conditions
and increased progressively during hypoxia to reach a maximum of 5.8 ± 0.53 mM (Fig.
2A,
P < 0.001).
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Fig. 2.
Plasma glucose ( ) and lactate ( ) concentrations
(A), glucose specific activity
(B), and glucose flux
(C) in rainbow trout under normoxic
(96% O2 saturation) and hypoxic
conditions (25% O2 saturation).
Shaded area indicates transition from normoxia to hypoxia. Values are
means ± SE (n = 8). dpm,
disintegrations per minute. * Significant differences from
normoxic values (P < 0.05).
C:  and  represent rates of
glucose appearance and disappearance, respectively, at the only time
when the animals were out of steady state. ** Rates of appearance
and disappearance were significantly different
(P < 0.05). At all other times
(), rates of appearance and disappearance were not different from
each other, and the values given are turnover rates.
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Hypoxia and glucose metabolism.
Glucose concentration, specific activity, and glucose flux throughout
the hypoxia experiments are plotted in Fig. 2. Normoxic values for
plasma glucose concentration and glucose turnover rate were 5.5 ± 0.7 mM and 5.4 ± 0.5 µmol · kg1 · min1,
respectively. Concentration was increased after 30 and 60 min of
hypoxia (P < 0.001), but returned to
a lower value that was not significantly different from normoxic
control after 90 min. Ra and
Rd values of glucose were only
different from each other 15 min after reaching 25% oxygen saturation
(P < 0.05). At that time,
Ra glucose was transiently higher
than baseline values. At all other times,
Ra and
Rd values were the same and not
significantly different from control values measured under normoxic
conditions.
Hypoxia and fatty acid metabolism.
Concentrations of individual NEFA and their percent contribution to
total plasma fatty acids before and after exposure to hypoxia are
presented in Table 1. Fatty acid
concentrations and their fractional contribution were not affected by
the change in water
PO2
(P > 0.05). Percent palmitate (16:0)
and percent docosahexaenoate (22:6) had the lowest coefficients of
variation of all individual fatty acids (11.9 and 10.1, respectively).
Figure 3 plots the effect of hypoxia on
plasma palmitate concentration, specific activity, and on the fractional contribution of this acid to total NEFA. Palmitate concentration remained at its normoxic level of 0.21 ± 0.02 mM throughout hypoxia (n = 7, P > 0.05). After separation of the different plasma lipids by TLC, 64.6 ± 0.03% of total
14C activity was found in the NEFA
fraction, and this percentage did not change over time
(P > 0.05). Palmitate specific
activity was increased during hypoxia
(P < 0.05), but percent palmitate was not affected by the change in oxygen availability
(P > 0.05). In normoxic fish, total
NEFA concentration and NEFA turnover rate averaged 0.98 ± 0.07 mM
and 5.9 ± 1.4 µmol · kg1 · min1
(n = 7), respectively (Fig.
4). Total NEFA concentration was not
affected by changes in water
PO2
(P > 0.05), but the turnover rate of
NEFA decreased significantly after 60 min of hypoxia and reached 53%
of normoxic levels at the end of the experiment
(P < 0.01).
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Table 1.
Plasma NEFA concentration and percent contribution to total NEFA
concentration in rainbow trout during normoxia and hypoxia
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Fig. 3.
Plasma palmitate concentration (A),
specific activity (B), and
fractional contribution to total nonesterified fatty acids (NEFA)
concentration (C) in rainbow trout
during the hypoxia experiments. Shaded area indicates transition from
normoxia to hypoxia. Values are means ± SE
(n = 7).
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Fig. 4.
Plasma NEFA concentration (A) and
turnover rate (B) in rainbow trout
during exposure to hypoxia. Values are means ± SE
(n = 7). * Significant
differences from normoxic levels (P < 0.05).
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Cold Exposure Experiments
Water temperature and
O2.
Changes in environmental temperature and in metabolic rate during cold
exposure are presented in Fig. 5. Mean
temperature was decreased from 15.0 to 5.8°C in 20 min, and the
water remained saturated with oxygen throughout the experiments. The
9°C change in temperature caused a progressive decrease in
O2
from 41.2 ± 2.3 to 21.7 ± 1.4 µmol
O2 · kg1 · min1
by the end of the experiment (P < 0.001). Observed activity levels remained minimal during the
measurements, except for ~1 min, when water temperature reached
10°C.
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Fig. 5.
Changes in water temperature during the cold exposure experiments
(line). Values are means ± SE (n = 8). Histogram shows
O2 in 15°C-acclimated
rainbow trout rapidly brought to 6°C. * Significant
differences from the 15°C control values
(P < 0.05).
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Cold exposure and glucose metabolism.
Glucose concentration, glucose specific activity, and glucose flux
throughout the cold exposure experiments are plotted in Fig.
6. Control values measured at 15°C for
plasma glucose concentration and turnover rate were 7.5 ± 1.1 mM
and 8.6 ± 1.3 µmol · kg1 · min1,
respectively. Glucose concentration was not significantly altered during the experiments, but specific activity was increased after 30 min at 6°C (P < 0.001).
Ra and
Rd values were never different from each other; therefore, the turnover rates presented in Fig. 6C were all calculated with the
steady-state equation of Steele (27). Cold exposure caused a large
decrease in glucose turnover rate to a mean value of 4.6 ± 0.7 µmol · kg1 · min1
(P < 0.001).
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Fig. 6.
Effects of cold exposure on plasma glucose concentration
(A), specific activity
(B), and turnover rate
(C) in 15°C-acclimated rainbow
trout. Shaded area indicates graded temperature decrease, and values
are means ± SE (n = 8).
* Significant differences from the 15°C control values
(P < 0.05).
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Cold exposure and fatty acid
metabolism. Concentrations of individual NEFA and their
percent contribution to total plasma fatty acids before and after cold
exposure are presented in Table 2. Fatty
acid concentrations were significantly increased after water
temperature reached 6°C (P < 0.05), but their fractional contributions to total NEFA were not
affected by this change (P > 0.05). Percent palmitate (16:0) and percent docosahexaenoate (22:6) had the lowest coefficients of variation of all individual fatty
acids (8.7 and 11.6, respectively). Palmitate concentration, specific
activity, and percent palmitate in total plasma NEFA throughout the
cold exposure experiments are plotted in Fig.
7. Under control conditions at 15°C,
palmitate concentration and percent palmitate averaged 0.18 ± 0.02 mM and 23.6 ± 0.1%. Acute exposure to 6°C caused a 40%
increase in palmitate concentration (P < 0.001), but had no effect on percent palmitate
(P = 0.25). After separation of the
different plasma lipids by TLC, 79.4 ± 0.7% of total
14C activity was found in the NEFA
fraction, and this percentage did not change over time
(P > 0.05). Palmitate specific
activity was increased 30 min after reaching 6°C and stayed
elevated until the end of the experiment
(P < 0.001). Total plasma NEFA
concentration went from a control value of 0.76 ± 0.09 to 1.07 ± 0.09 mM after 2 h at 6°C (P < 0.0001, Fig.
8A). The
turnover rate of plasma NEFA averaged 4.8 ± 0.7 µmol · kg1 · min1
under 15°C control conditions, and it was significantly decreased after exposure to 6°C (P < 0.001, Fig. 8B).
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Table 2.
Plasma NEFA concentration and percent contribution to total NEFA
concentration at 15°C and after 90 min at 6°C in rainbow trout
acclimated to 15°C
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Fig. 7.
Effects of cold exposure on plasma palmitate concentration
(A), specific activity
(B), and fractional contribution to
total NEFA concentration (C)
in 15°C-acclimated rainbow trout. Shaded area indicates
graded temperature decrease, and values are means ± SE
(n = 8). * Significant
differences from the 15°C control values
(P < 0.05).
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Fig. 8.
Effects of cold exposure on plasma NEFA concentration
(A) and turnover rate
(B) in 15°C-acclimated rainbow trout. Shaded area
indicates graded temperature decrease, and values are means ± SE
(n = 8). * Significant
differences from the 15°C control values
(P < 0.05).
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Q10 effects on
O2
and fluxes of oxidative fuels.
Changes in metabolic rate, glucose flux, and fatty acid flux during
cold exposure are presented in Fig. 9 as a
percent of the 15°C control values. The three parameters responded
similarly to the 9°C drop in water temperature and their
Q10 values were not different from
each other (P > 0.05). Metabolic
rate, glucose turnover, and fatty acid turnover had
Q10 values of 2.2 ± 0.2, 1.9 ± 0.1, and 2.1 ± 0.1, respectively.
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Fig. 9.
Relative effects of cold exposure on
O2 (), glucose turnover
rate ( ), and NEFA turnover rate () in 15°C-acclimated rainbow
trout. Values are percentages of the control 15°C rates (dotted
line) and are means ± SE (n = 8).
Shaded area indicates graded temperature decrease. Turnover rates of
glucose and NEFA were not different from each other
(P > 0.05).
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DISCUSSION |
Effects of hypoxia
Under severe hypoxia, rainbow trout are able to maintain normoxic
metabolic rates (Fig. 1) (24) by increasing oxygen extraction through
neuronal and hormonal mechanisms (9, 22). Anaerobic glycolysis is
stimulated, as indicated by a steady accumulation of lactate in plasma
(Fig. 2A) (2, 7, 8), and hypoxia is
thought to produce a general shift in oxidative fuel preference, from
predominantly lipids and proteins to carbohydrates (30, 31). Here, we
report the first fatty acid turnover rates in fish and show that the
importance of this fuel decreases throughout hypoxia (Fig.
4B). At the same time, hepatic
glucose production is briefly stimulated at the onset of the hypoxic
stress, but this increase is not accompanied by a change in glucose
utilization (Fig. 2C), and a
temporary imbalance between the rates of production and utilization
leads to hyperglycemia (Fig. 2A).
This change in blood glucose concentration was also noticed in the only
other study where glucose turnover rate was measured in hypoxic fish (8). However, the transient increase in glucose production could not be
detected at the time, because bolus injection was used and this
technique does not allow continuous measurements under non-steady-state
conditions (12). Severe hypoxia is known to produce the release of
catecholamines and other stress hormones that are probably responsible
for the hyperglycemic response through stimulation of glycogenolysis
and/or gluconeogenesis (31, 34, 41).
The lower NEFA flux and the steady NEFA concentration observed in
hypoxic fish can be explained by a parallel decrease in the rates of
NEFA oxidation and mobilization. The actual signal for lowering NEFA
flux may have been high plasma glucose concentration. It was suggested
long ago that hyperglycemia could inhibit NEFA mobilization
independently from hormonal control (25). More recently, this
inhibition of lipolysis has been demonstrated in humans (4, 40), and
our results suggest that the same reciprocal interaction between plasma
glucose concentration and fatty acid mobilization is also present in
fish. However, the Ra of glycerol was not quantified in our experiments, and, consequently, the possibility that the decrease in NEFA flux was caused by a stimulation of reesterification cannot be totally ruled out. Future development of
adequate techniques to quantify the turnover rate of glycerol in fish
will be necessary to measure reesterification and to resolve this
issue. Finally, the fact that glucose production returns to normoxic
levels after 1 h of hypoxia suggests that the observed increase in
anaerobic glycolysis is mainly fueled by glycogen reserves rather than
by circulating glucose.
Effects of Temperature
Compensatory mechanisms associated with temperature acclimation have
been investigated thoroughly (5, 10, 13, 17), but the effects of acute
temperature changes have not received much attention. This study
quantifies the impact of a rapid temperature change on the metabolic
fuel kinetics of rainbow trout. A 9°C drop in water temperature
causes a 50% decrease in
O2
(Fig. 5), and this decrease is accompanied by equivalent changes in the
turnover rates of glucose (Fig. 6C)
and NEFA (Fig. 8B). No preferential
inhibition of glucose or fatty acid mobilization is taking place when
water temperature decreases (Fig. 9) and, therefore,
Q10 values for the fluxes of both
substrates as well as oxygen are similar, ranging from 1.9 to 2.2. Identical Q10 values have also
been reported for trout
O2
after temperature acclimation (3), and this suggests that glucose and
NEFA fluxes would remain at the low levels measured at the end of our
experiments, even after complete acclimation to 6°C. The changes in
substrate turnover rates reported here probably reflect how rates of
substrate oxidation are affected. However, further studies will be
necessary to quantify fuel oxidation directly, thereby providing a
clearer understanding of the effects of temperature on energy
metabolism.
Cold exposure elicited a large increase in plasma NEFA concentration
(Fig. 8A), but the difference
between the rates of fatty acid release and uptake responsible for it
was too small to be detectable, and we can only speculate on the
mechanisms involved in this change. We can say with certainty that the
rapid decrease in water temperature did not have exactly the same
effect on release and uptake. The resulting imbalance may have been
caused by differential temperature effects on membrane fluidity or on
the capacity of various lipid-binding proteins to transport fatty
acids.
Relationship Between Substrate Concentration and Flux
This study shows that variation in plasma metabolite levels do not
reflect changes in flux, and, therefore, that concentration cannot be
used to speculate on potential dynamic changes in flux. The relative
effects of hypoxia and low temperature on the concentrations and fluxes
of glucose and NEFA are summarized in Fig.
10. Concentration and flux never changed
in parallel during our experiments, and it would have been impossible
to predict the effects of environmental changes on substrate fluxes
simply by examining plasma metabolite concentrations. Hypoxia caused a
large increase in glucose concentration, but had no effect on glucose
flux; in contrast, it did not increase NEFA concentration
significantly, but caused a sharp decrease in NEFA flux. Lowering water
temperature caused a parallel decrease in glucose and NEFA fluxes, but
glucose concentration was maintained steady, whereas NEFA concentration
increased by more than 40%. The experimental manipulations performed
in this study illustrate clearly that flux and concentration can change
independently. For example, it is known from the mammalian literature
that plasma NEFA concentration and flux are strongly correlated during
exercise (11, 21), and, therefore, NEFA concentration is commonly used as an index of the rate of NEFA utilization in this context. Present results warn us that such a correlation is only valid under specific circumstances and should never be viewed as a universal phenomenon. In
our experiments, and contrary to expectation, a strong increase in NEFA
concentration was accompanied by a large decrease in NEFA flux when
water temperature was lowered. In future studies, metabolite fluxes
will have to be measured directly to investigate the impact of
environmental, hormonal, and other changes on fish metabolism. Furthermore, published findings from experiments where plasma concentrations were used to draw conclusions about fluxes should be
reevaluated with the above concerns in mind (e.g., see Refs. 20, 23,
32).
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Fig. 10.
Relative effects of acute hypoxia and cold exposure on plasma
concentrations (open bars) and fluxes (solid bars) of oxidative fuels
in rainbow trout.
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Plasma NEFA Composition and Use of Palmitate as a Tracer
The fractional contribution of individual fatty acids to total plasma
NEFA was not affected by exposure to hypoxia (Table 1) or low
temperature (Table 2). Palmitate (16:0) and docosahexaenoate (22:6)
were the most abundant fatty acids in rainbow trout plasma as reported
previously (31). Palmitate is commonly used to measure NEFA kinetics in
mammals because its fractional contribution to total NEFA shows the
lowest coefficient of variation in this group of vertebrates (19). In
trout, we found that 16:0 and 22:6 had the lowest coefficients among
all circulating fatty acids and, therefore, that these two lipids are
the best representatives of changes in total NEFA concentration. This
suggests that labeled 16:0 and 22:6 are appropriate tracers to measure
NEFA kinetics in rainbow trout. In the present study, we have
selected 16:0 because it is more readily available commercially, but
also because we were more specifically interested in energy metabolism,
and 16:0 is more likely to be used as an oxidative fuel than 22:6 (33).
However, polyunsaturated fatty acids (PUFA) are known to play an
important role in the reorganization of phospholipids to maintain
adequate membrane fluidity during temperature changes. The substitution
of saturated with unsaturated fatty acids in membrane phospholipids is
known to occur rapidly in fish exposed to cold water (16, 35).
Consequently, the changes in NEFA turnover rates measured in our cold
exposure experiments with palmitate as a tracer may have underestimated
temperature effects on PUFA fluxes.
Conclusions
This study provides the first continuous measurements of oxidative
substrate kinetics in fish exposed to a rapid decrease in oxygen
availability or in temperature. It shows that acute hypoxia causes a
progressive decrease in the turnover rate of NEFA, together with a
transient increase in hepatic glucose production, whereas a rapid drop
in water temperature induces strong and equivalent declines in glucose,
NEFA, and oxygen fluxes. Our results also demonstrate that the
fluctuations in glucose and NEFA fluxes caused by environmental
stresses are not accompanied by interpretable changes in the
concentrations of these metabolites in plasma.
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ACKNOWLEDGEMENTS |
F. Haman was the recipient of an Ontario Graduate Scholarship, and
this work was supported by a National Sciences and Engineering Research
Council Research Grant (Canada) to J.-M. Weber.
| |
FOOTNOTES |
Address for reprint requests: J.-M. Weber, Biology Dept., Univ. of
Ottawa, 30 Marie Curie, Ottawa, Ontario, Canada K1N 6N5.
Received 2 June 1997; accepted in final form 3 September 1997.
| |
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