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First published online November 19, 2007
Journal of Experimental Biology 210, 4254-4261 (2007)
Published by The Company of Biologists 2007
doi: 10.1242/jeb.005835
Temperature effects on metabolic rate of juvenile Pacific bluefin tuna Thunnus orientalis
1 Hopkins Marine Station, Stanford University, Pacific Grove, CA 93950,
USA
2 Monterey Bay Aquarium, Monterey, CA 93940, USA
* Author for correspondence present address: Ecology and Evolutionary Biology, 321 Steinhaus Hall, University of California, Irvine, CA 92697-2525, USA (e-mail: jblank{at}uci.edu)
Accepted 19 September 2007
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Summary |
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O2) was
measured at ambient temperatures of 8–25°C and swimming speeds of
0.75–1.75 body lengths (BL) s–1. Pacific
bluefin swimming at 1 BL s–1 per second exhibited a
U-shaped curve of metabolic rate vs ambient temperature, with a
thermal minimum zone between 15°C to 20°C. Minimum
O2 of
175±29 mg kg–1 h–1 was recorded at
15°C, while both cold and warm temperatures resulted in increased
metabolic rates of 331±62 mg kg–1 h–1
at 8°C and 256±19 mg kg–1 h–1 at
25°C. Tailbeat frequencies were negatively correlated with ambient
temperature. Additional experiments indicated that the increase in
O2 at low
temperature occurred only at low swimming speeds. Ambient water temperature
data from electronic tags implanted in wild fish indicate that Pacific bluefin
of similar size to the experimental fish used in the swim tunnel spend most of
their time in ambient temperatures in the metabolic thermal minimum zone.
Key words: endothermy, metabolic rate, temperature, thermoregulation, tuna
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Introduction |
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TOPSummaryIntroductionMaterials and methodsResultsDiscussionReferences |
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;
Block et al., 1993;
Brill and Bushnell, 2001).
Tunas have evolved vascular countercurrent heat exchangers, retia
mirabilia, to conserve metabolic heat and maintain elevated temperatures
in slow-twitch muscles, viscera, eyes and brain
(Carey and Teal, 1966;
Linthicum and Carey, 1972).
Substantial thermal gradients are often present within the body of an
individual fish (Carey and Teal,
1966) while some tissues, including the heart and gills, operate
at ambient water temperature. Data from electronic tags indicate that Pacific
bluefin (Thunnus orientalis) and yellowfin tuna (T.
albacares) range through a thermally variable environment
(Block et al., 2001; Block et
al., 1997; Kitagawa et al.,
2007; Marcinek,
2000; Schaefer et al.,
2007), resulting in fluctuating tissue temperatures that are
affected by species, body size, ambient water temperature, feeding status and
activity level. The relationships of the metabolic rates of tunas to ambient
temperature are potentially complex and may be distinct from those of
ectothermic fish or terrestrial endotherms.
Measurements of oxygen consumption rates
(O2) in tunas
are complicated by their large size and need to swim continuously to ventilate
the gills and generate hydrodynamic lift
(Magnuson, 1973). To date,
most studies of tuna metabolism have examined tropical or warm temperate
species, and only a few studies have examined acute temperature changes
(Korsmeyer and Dewar, 2001).
Standard metabolic rates of 250-500 mg O2 kg–1
h–1 have been measured at 25°C in 0.5–4 kg kawakawa
(Euthynnus affinis), yellowfin, and skipjack tuna (Katsuwonus
pelamis) (Boggs and Kitchell,
1991; Brill, 1979;
Brill, 1987;
Bushnell and Brill, 1992;
Dewar and Graham, 1994;
Gooding et al., 1981;
Korsmeyer et al., 1997). In
swimming yellowfin tuna, acute temperature changes between 18 and 30°C
yielded a mean Q10 of 1.67 for
O2
(Dewar and Graham, 1994).
Anesthetized yellowfin, skipjack and kawakawa
O2 exhibited
Q10 values of 2.3, 2.4 and 3.2, respectively between 20 and
25°C (Brill, 1987). While
these Q10 values are similar to those of ectothermic teleosts,
there has been little research on the relationship of metabolic rate to
ambient water temperature in the cold–temperate tunas such as albacore
(Thunnus alalunga) or bluefin tunas (T. thynnus, T.
orientalis and T. maccoyii). Shipboard measurements of albacore
metabolic rates have been limited to a 3°C temperature range or less
(Graham and Laurs, 1982;
Graham et al., 1989).
Recent measurements of
O2 in swimming
7–10 kg Pacific bluefin tuna indicate that Pacific bluefin have higher
metabolic rates than yellowfin tuna of similar size at 20°C
(Blank et al., 2007). Minimal
metabolic rates of fasted bluefin and yellowfin tunas in the swim tunnel were
222±24 mg O2 kg–1 h–1 and
162±19 mg O2 kg–1 h–1,
respectively, at 20°C. Rates up to 498±55 mg O2
kg–1 h–1 were recorded in bluefin at 1.8
BL s–1. Routine metabolic rate in southern bluefin
tuna (Thunnus maccoyii) swimming freely in a circular mesocosm was
recently measured and reported as mean metabolic rate of 460 mg O2
kg–1 h–1 for fasted 20 kg fish in 19°C
water (Fitzgibbon et al.,
2006). Feeding resulted in elevated
O2 for a period
of 20–45 h accompanied by increases in voluntary swimming speed
(Fitzgibbon et al., 2007).
Adult Atlantic bluefin tuna maintain large elevations in muscle temperature
(Carey and Teal, 1969) and
juvenile Pacific bluefin (12–20 kg) have the capacity to maintain
relatively stable muscle temperatures 6–8°C above ambient water
temperatures (Marcinek et al.,
2001). Visceral temperatures of wild Pacific bluefin exhibit diel
cycles of warming associated with specific dynamic action, resulting in
elevations of 4–12°C above ambient water temperature for 6–24
h following feeding (Kitagawa et al.,
2007). In contrast to the warm muscle and viscera of swimming
bluefin tunas, the heart remains at or near ambient water temperature in all
tuna species (Brill et al.,
1994; Carey et al.,
1984), resulting in pronounced effects of ambient temperature on
heart function (Blank et al.,
2002; Blank et al.,
2004; Korsmeyer et al.,
1997; Landeira-Fernandez et
al., 2004).
The availability of Pacific bluefin tuna in captivity and the ability to maintain tunas in the swim tunnel for extended periods of up to 6 days provides an unprecedented opportunity to study the response of Pacific bluefin tuna metabolic rates to changes of ambient water temperatures and swimming speeds. Measurements at 8–25°C indicate that metabolic rates of bluefin tuna swimming at low speed reach a thermal minimum zone (TMZ) that corresponds to the preferred sea surface temperatures of similar sized Pacific bluefin tunas in the wild.
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). Fish were
held for 58–162 days prior to use in swim tunnel experiments and
archival tags were implanted in the visceral cavity during this period. Mean
curved fork length (CFL) of 9 bluefin used in the swim tunnel was
76±4 cm (range 70–84 cm), and mass was 8.8±0.8 kg (range
7.4–9.9 kg). All fish were feeding prior to experiments. Food was
withheld from the entire tank for 45–72 h prior to the introduction of a
fish into the swim tunnel.
Swim tunnel
Bluefin tuna swam in a swim tunnel consisting of an 870 l acrylic
respirometer chamber with a square 45 cmx45 cmx135 cm working
section contained in a 1500 l external tank for thermal insulation (Loligo
Systems, Denmark). Turbulence was minimized by two honeycomb sections upstream
of the working section and additional flow straighteners. The sides of the
working section were marked with vertical stripes of black tape at 10 cm
intervals to assist the fish in maintaining position in the tank. Water
velocities were calibrated by video analysis of dye injections and corrected
for solid blocking effects for each fish
(Bell and Terhune, 1970). Fish
cross-sectional area ranged from 6 to 10% of working section area and blocking
correction ranged from 5 to 11% of uncorrected velocity. Seawater was supplied
to the respirometer from a 20 000 l reservoir, which was continuously filtered
and aerated. Temperature of the swim tunnel was adjusted to the target
temperature ±0.1°C by a 6800 W heat pump and addition of warm or
cold filtered seawater to the 20 000 l reservoir. The entire swim tunnel was
enclosed by black plastic sheeting to minimize light fluctuations and access
to the building was limited while the experiment was in progress to minimize
disturbance. A video camera and a mirror mounted at a 45° angle above the
working section provided an overhead view of the swimming fish, allowing
continuous monitoring of its swimming behavior.
Introduction of the fish
Each tuna was captured in a water-filled nylon sling and a measurement of
its CFL was taken. The sling holding the fish was lifted from the
holding tank and carried to the swim tunnel. The front end of the sling was
submerged in seawater flowing at 
1 BL s–1 in
the working section and opened. One researcher manually guided the fish from
the sling into the flowing water, while the other researcher removed the
sling. The lid of the working section was used to cover the front of the
working section as the fish was introduced and then moved to cover the rest of
the working section. One researcher positioned a hand at the rear of the
working section to prevent the fish from contacting the rear grate as it
adjusted to the swim tunnel. Once the fish was swimming steadily,
approximately 30–45 min after introduction, the lid of the working
section was secured with wing nuts and the pumps supplying aerated seawater to
the swim tunnel were set to alternate between flushing the swim tunnel with
aerated seawater and closing for measurements of
O2. Fish that
did not swim steadily were removed from the swim tunnel and returned to the
holding tank.
Respirometry procedures
Oxygen consumption was measured by stopped-flow respirometry as detailed by
Steffensen (Steffensen, 1989).
The swim tunnel was closed for 10 min for measurement of
O2 and then
flushed with aerated seawater for 10 min to restore dissolved oxygen
(DO2). These steps were repeated throughout the experiment.
DO2 remained above 80% of air saturation levels throughout each
experiment and was generally above 90% saturation except at 25°C.
Temperature and oxygen content of the seawater in the swim tunnel were logged
at 10 or 15 s intervals by a temperature-compensated multiprobe (Yellow
Springs Instruments Model 556, Yellow Springs, OH, USA) and
O2 was
calculated from the rate of decline in DO2 during each closed
period. The O2 electrode was calibrated in air-saturated seawater
at 20°C prior to each experiment, and the calibration was found to drift
by less than 1% over the duration of each experiment.
Series I
Six bluefin tuna (mass=9.1±0.6 kg, range 8.1–9.9 kg) were used
in the initial series of measurements. Three of these fish were previously
used in separate experiments on different dates
(Blank et al., 2007) and were
reintroduced to the swim tunnel for this study. Once metabolic rate
measurements were initiated, the swim speed was adjusted to 1.0 BL
s–1 and the fish was allowed to acclimate to the respirometer
overnight while temperature was maintained at 20°C, matching the
acclimation temperature in the holding tank. Following the initial acclimation
period, or following speed tests conducted in three individuals according to
the previously described protocol (Blank et
al., 2007), a series of at least 12 consecutive
O2 measurements
was completed at 20°C. Temperature was then adjusted and at least 12
measurements were taken at 15°C, followed by 10°C, 8°C and
25°C (Fig. 1). Attempts to
measure O2 at
temperatures below 8°C were unsuccessful, as some fish stopped swimming or
began to lose righting responses when the swim tunnel was cooled further. Each
test temperature was held constant for a minimum of 4 h throughout the
measurement and flush cycles. If the behavior or
O2 of the fish
was irregular, this period was extended to increase the number of measurements
taken.
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O2 measurements.
In some experiments, a 12 h:12 h light:dark cycle (with dim light at night)
was maintained, matching the cycle in the holding tanks. In most cases, the
lighting changes appeared to startle the fish and constant dim light was used
instead. Control measurements at 20°C were repeated at or near the end of
the experiment to confirm the repeatability of the baseline metabolic rate.
O2 at 20°C
changed by –4 to +17% (mean=9%) over the period of the experiment. When
all measurements were completed, the fish was removed from the swim tunnel and
weighed on a cushioned vinyl V-board with a damp cloth over its eyes. The
length of the fish was verified and the fish was returned to its holding tank.
After each experiment, the swim tunnel was sealed and background respiration
measured. In all cases, background respiration was negligible.
Series II
In a second series of experiments on a different group of three fish
(mass=8.1±0.6 kg, range 7.4–8.4 kg), the effects of swimming
speed on O2 were
compared at three different temperatures. Each fish was introduced to the swim
tunnel as described above and allowed to acclimate to the swim tunnel for
2–3 h while swimming at 1.0 BL s–1 at
20°C. The fish was then presented with a practice series of speed changes
in which speed was increased in 0.15 BL s–1
increments during each flush period up to a maximum of 1.75 BL
s–1. Following completion of this `practice speed test', the
speed was reduced to 1.0 BL s–1 and the fish was
allowed to acclimate to the swim tunnel overnight for a minimum of 15 h. On
the following day, speed was elevated or lowered in increments of 0.10 or 0.15
BL s–1 and a series of at least four consecutive
O2 measurements
was completed at each speed setting from 0.75 BL s–1
to 1.75 BL s–1. Fish were observed to swim closer to
the back of the swim tunnel at faster speeds, so speeds above 1.75 BL
s–1 were not tested to minimize the risk of damage to the
caudal fin of the fish. No attempt was made to measure maximum swimming speed
or O2max. After
the initial speed test, the temperature of the swim tunnel was adjusted to 8
or 25°C, the fish was allowed to equilibrate for at least 4 h, and the
entire speed test was repeated at the new temperature. Speed tests were
conducted at each temperature on successive days and control measurements were
taken at 20°C before the fish was removed from the swim tunnel, as
described above.
Data analysis
While in the swim tunnel, the fish was monitored continuously via
closed circuit video and any aberrant swimming behaviors or external
disturbances were noted. Individual measurements of
O2 and other
variables associated with disturbances such as earthquakes, power outages, air
bubbles in the respirometer and spontaneous attempts by the fish to turn
around in the working section were excluded from further analysis. The first
60 min of data following each ambient temperature change were also excluded to
allow for thermal equilibration. The mean of all remaining measurements at a
given speed and/or temperature was then taken as the
O2 for that
condition for subsequent calculations and statistical analysis (minimum
N=5).
Tailbeat frequency was measured by an observer watching the live video
display of the overhead view of the swimming fish. Sixty tailbeats were timed
with a stopwatch 3 times and the computed tailbeats min–1
from the three counts were averaged. This process was repeated at least two
times for each fish at each speed. Gross cost of transport (GCOT) was
calculated from
O2 and swimming
speed using an oxycalorific coefficient of 14.1 J mg
O2–1 for a mixed diet
(Videler, 1993). For series I,
data collected at different temperatures were compared by one-way analysis of
variance (ANOVA) using Systat 11.0. Post-hoc comparisons were made
with Tukey's HSD and significance was assessed at P<0.05. For
series II, two-way ANOVA was used to assess the combined effects of
temperature and swimming speed on
O2, tailbeat
frequency and GCOT. GCOT data were log-transformed to assure normality. Data
are presented as means ± s.d.
Archival tagging
Two models of archival tags (Lotek LTD 1400 and LTD 2310, Lotek Wireless,
Inc., Newmarket, Ontario, Canada) were implanted in fish to record visceral
temperatures at 4 or 8 s intervals as the fish swam in the respirometer. For
tag implantation, the water level in the holding tank was lowered to 90 cm and
a bluefin tuna was gently guided into a water-filled nylon sling. The fish was
placed ventral-side up, its eyes covered with a moist cloth, and its gills
irrigated with a hose placed in its mouth. A sterile #22 surgical blade was
used to make a small incision in the ventral body wall and the tag was
inserted in the peritoneal cavity with the sensor stalk protruding from the
ventral body wall. The incision was sutured shut and the fish was released to
recover in the tank for a minimum of 7 days. Fish were observed to feed as
soon as 2 days after tag implantation and all tagged fish had recovered prior
to use in swim tunnel experiments. Tags were programmed to record visceral
temperature (both tags) and/or ambient water temperature (LTD 2310 only) at
intervals of 4 or 8 s. An additional Lotek 2310 tag was synchronized and
placed in the swim tunnel to record ambient temperature. All fish were tagged,
and tags were recovered from three fish used in Series I experiments
post-mortem and time series data were downloaded. Each of the three
tags was embedded in the visceral mass in contact with the pyloric caecum. The
mean visceral and ambient temperatures during the last hour at each ambient
test temperature were considered to represent steady state and used for
analysis of thermal excess.
Wild bluefin (N=10, CFL=75.2±2.8 cm) were tagged
offshore of San Diego, California using previously published procedures
(Block et al., 1998;
Kitagawa et al., 2007). Tags
were programmed to collect pressure, light and temperature data every 4, 8, 16
or 32 s. Since the LTD2310 tags were implanted in the peritoneal cavity of the
tuna, recapture of the fish and recovery of the tag were necessary in order to
recover the collected data. The tagged bluefin tuna were recaptured after
241±131 days at liberty and data was downloaded from the tags. For each
day, we extracted pressure, ambient water temperature, and body temperature
data.
All surgical, tagging and other experimental procedures were conducted in accordance with Stanford University institutional animal use protocols.
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O2 values
ranging from 175±29 mg kg–1 h–1 to
331±62 mg kg–1 h–1
(Fig. 2A). An apparent thermal
minimum zone was reached between 15 and 20°C with similar
O2
(P=0.95, 15°C vs 20°C) and a Q10 of only
1.25 in this temperature range. Bluefin tuna swimming at a constant speed of
1.0 BL s–1 increased
O2 significantly
from 175±29 mg kg–1 h–1 at 15°C
to a maximum of 331±62 mg kg–1 h–1 at
8°C (P<0.001), resulting in a mean Q10 of 0.41
between 8 and 15°C. Tailbeat frequency increased from 103±5
tailbeats min–1 at 25°C to 127±11 tailbeats
min–1 at 8°C (Fig.
2B, P=0.006). At 8–10°C, swimming behavior
became noticeably erratic. Bluefin periodically made contact with the front
honeycomb or front corner of the swim tunnel's working section and produced
bursts of rapid tailbeats. From 20–25°C, bluefin tuna
O2 increased
from 193±25 to 256±19 mg kg–1
h–1 (Q10 of 1.8;
Fig. 2A).
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O2 at each swim
speed. At both 20 and 25°C, increasing speed from 1.0 to 1.75 BL
s–1 resulted in a 1.7-fold increase in
O2
(P<0.001). In contrast, bluefin tuna cooled to 8°C showed no
change of O2
with swim speed (P=0.89). A significant interaction of temperature
and swim speed was detected (P=0.015). Tailbeat frequencies recorded
at 8°C were higher than those at 20 and 25°C (P<0.001,
Fig. 3B). GCOT calculated from
O2 at each speed
showed a significant interaction of speed and temperature (P=0.023),
with a maximum GCOT of 2.8±0.5 J kg–1
m–1 at 0.75 BL s–1 and 8°C
(Fig. 4).
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Discussion |
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O2 to ambient
temperature differs from the thermal response of most fishes, and more closely
resembles the response of an endothermic animal to ambient temperature.
Previous studies of metabolic rate in yellowfin and skipjack tunas have
reported Q10 values of 1.7–3.2
(Brill, 1987;
Dewar and Graham, 1994), which
are within the typical range for ectothermic fish. However, these studies used
small individuals of tropical tuna species at ambient temperatures that were
no lower than 18°C. Tunas in this size range are capable of elevating
muscle temperature above ambient (Dewar et
al., 1994; Dickson,
1994), but maintain lower thermal excess temperatures than larger
individuals and may be less capable of thermoregulation. Thus, they might
exhibit a qualitatively different metabolic response to ambient temperature
change than the Pacific bluefin in this study.
The observation that bluefin tuna metabolic rates are minimized at intermediate temperatures and increase at colder temperatures is unique among teleost fishes. Several potential explanations for this pattern must be considered, including stress related to confinement in the swim tunnel at low temperature, failure or decreased efficiency of the slow-twitch muscle at low temperature, and metabolic or behavioral thermoregulation.
High metabolic rates and tailbeat frequencies at low temperatures might
indicate a stress or avoidance response as the fish encounter an unfamiliar or
unpleasant ambient water temperature. The transient elevation of
O2 following a
power outage (Fig. 1)
illustrates the potential for stress to elevate recorded
O2 values
(Brett, 1962;
Steffensen, 1989).
Observations of fish pushing against the front of the working section while
producing bursts of rapid tailbeats at low ambient water temperatures
(8–10°C) could be construed to reflect stress or escape behavior,
but the possible role of stress at low temperatures remains unclear.
The increase in
O2 at low
temperatures in Series I may be associated with a cold-induced decline of
power from slow-twitch muscle, resulting in earlier recruitment of fast-twitch
muscle (Rome et al., 1984) and
a potential increase in locomotor cost as the fish swam at a constant speed.
Optimum frequency and power output of yellowfin tuna slow-twitch muscle are
temperature dependent (Altringham and
Block, 1997) and slow-twitch muscle from the endothermic salmon
shark is unable to produce positive work at low temperatures
(Bernal et al., 2005), raising
the possibility that bluefin tuna slow-twitch muscle would be similarly
impaired at cold temperatures. The decline in optimum contraction frequency at
low temperature in vitro indicates that cold muscle should be least
effective at high swimming speeds
(Altringham and Block, 1997).
However, cold ambient temperatures increased
O2 and GCOT only
at low speeds (Fig. 3A,
Fig. 4), suggesting that muscle
power output was not limiting to swimming capacity at low temperature.
Alternatively, elevated
O2 at low
ambient temperature may reflect metabolic or behavioral thermoregulation.
Visceral thermal excess was less than 2°C at all ambient temperatures
(Fig. 5A,B), reflecting the
45–72 h fasting period preceding introduction of the fish to the swim
tunnel. This observation of a low visceral thermal excess in fasted bluefin
tuna is in accord with large data sets from wild fish
(Itoh et al., 2003;
Kitagawa et al., 2001). Muscle
temperature was not measured in this study to avoid potential injury
associated with implantation of temperature loggers in the muscle. Therefore,
regulation of muscle temperature cannot be ruled out. The bursts of rapid
tailbeats and increased mean tailbeat frequencies recorded at low temperatures
(Fig. 2B,
Fig. 3B) provide a potential
mechanism for increased metabolic heat production in the swimming muscle. A
similar response was observed in restrained albacore tuna, which increased
tailbeat frequency by 20% and reduced tailbeat amplitude in association with
thermoregulation at ambient temperatures below 14°C
(Graham and Dickson, 1981).
Albacore were able to maintain stable muscle temperatures during more than 2 h
of exposure to low ambient temperatures.
Experiments in which temperatures are recorded in the slow-twitch muscle of
bluefin during respirometry would be required to clarify whether maintenance
of muscle temperature might be related to elevation of
O2 at low
temperatures in swimming bluefin. Acoustic tagging data from wild fish
indicate that slow-twitch muscle temperatures are more stable than visceral
temperatures in both large and small Atlantic and Pacific bluefin
(Carey and Lawson, 1973;
Carey and Teal, 1969;
Marcinek et al., 2001).
However, it is unclear whether 8–10 kg bluefin are capable of
maintaining stable muscle temperatures during extended exposure to
8–10°C water.
Importantly, the 15–20°C range at which
O2 is minimized
in the swim tunnel corresponds well to the modal ambient temperatures recorded
in archival tag data from similar sized wild 8–10 kg Pacific bluefin
(Fig. 5)
(Kitagawa et al., 2006). These
results suggest that juvenile fish may occupy regions with ambient water
temperatures that result in the lowest metabolic costs. Small fish tracked off
the west coast of North America sometimes experience lower sea surface
temperatures (12–14°C) for prolonged periods
(Kitagawa et al., 2007), which
might entail higher energetic costs. However, numerous factors, such as
thermal acclimation to colder waters, faster swimming speeds in the wild, and
specific dynamic action could conceivably shift the thermal optimum in wild
fish.
Based on electronic tag data, 8–10 kg Pacific bluefin such as those
in this study make occasional dives into 8–10°C water, but do not
spend long periods at these temperatures. Archival tag records from both adult
and juvenile Atlantic and Pacific bluefin tuna frequently show `bounce' diving
patterns in which the fish alternate diving into cool waters below the
thermocline and returning to the warmer surface waters, typically on a time
scale of minutes (Blank et al.,
2004; Gunn and Block,
2001; Kitagawa et al.,
2001). Bluefin encountering low temperatures at depth might adjust
dive duration to maintain skeletal, cardiac muscle and/or visceral
temperatures above a threshold for adequate tissue function
(Blank et al., 2004). As diving
below the thermocline is often associated with foraging, burst swimming
activity and specific dynamic action may also contribute to metabolic heat
production during dives. How metabolic rate, body temperature and diving
behavior are interrelated has yet to be fully explored. Instrumentation of
bluefin for muscle temperature measurement and chronic blood sampling during
swim tunnel respirometry at varying ambient temperatures would provide
valuable information on the physiological factors determining performance
limits of bluefin tuna at high and low ambient temperatures.
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Acknowledgments |
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References |
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TOPSummaryIntroductionMaterials and methodsResultsDiscussionReferences |
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Significant difference from TBF at 8°C.
