Effects of temperature on intracellular [Ca2+] in trout atrial myocytes
1 Simon Fraser University, Biological Sciences, Burnaby, British Columbia,
V5A 1S6, Canada
2 University of Joensuu, Department of Biology, PO Box 111, 80101 Joensuu,
Finland
* Author for correspondence at present address: School of Biomedical Sciences, University of Leeds, Leeds, LS2 9JT, UK (e-mail: hollys{at}sfu.ca)
Accepted 21 August 2002
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Summary |
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Key words: L-type Ca2+ current (ICa), calcium transient, sarcoplasmic reticulum, action potential, frequency, fish hearts, temperature, rainbow trout, Oncorhynchus mykiss
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Introduction |
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In mammals, the temperature sensitivity of extracellular and intracellular
Ca2+ fluxes are well characterized. Transsarcolemmal
Ca2+ influx via the L-type Ca2+ channel current
(ICa) is strongly influenced by temperature, with a Q10
ranging from 2 to 3.5 (Cavalie et al.,
1985; Herve et al.,
1992
). Cold temperatures reduce SR function in many mammalian
hearts, impairing both release (Bers,
1987
; Sitsapesan et al.,
1991
) and uptake (Liu et al.,
1991
,
1997
;
Wang et al., 2000
) of SR
Ca2+ stores. Under most conditions, excitationcontraction
coupling in rainbow trout heart is thought to be driven primarily by
extracellular Ca2+ influx via ICa, with SR
Ca2+ release playing a secondary role
(Tibbits et al., 1992
;
Hove-Madsen, 1992
;
Keen et al., 1994
;
Shiels and Farrell, 1997
).
Recent studies of ICa in trout myocytes indicate that the
temperature sensitivity of peak current amplitude is not very different from
that of mammals (Q10 of approximately 1.7-2.1;
Kim et al., 2000
;
Shiels et al., 2000
) but that
the total transsarcolemmal Ca2+ influx is fairly well maintained
during temperature change (Shiels et al.,
2000
). Furthermore, the fish cardiac SR has been shown to uptake
and release Ca2+ at temperatures well below those known to inhibit
SR function in most mammals (Aho and
Vornanen, 1999
; Hove-Madsen et
al., 2001
; Shiels et al.,
2002a
; Tiitu and Vornanen,
2002
).
Despite the fact that the individual Ca2+ flux pathways in trout
hearts seem to demonstrate some tolerance to temperature change, in
situ heart and in vitro cardiac muscle strip studies show both
inotropic and chronotropic responses to temperature change
(Keen et al., 1994;
Shiels and Farrell, 1997
;
Aho and Vornanen, 1999
). We
wanted to understand to what extent these responses might be mediated by
changes in cellular Ca2+. The transient change in
[Ca2+]i during excitationcontraction coupling
represents the sum of both extracellular and intracellular Ca2+
flux, and its amplitude is directly related to the strength of cardiac
contraction (Yue, 1987
). Thus,
the focus of the present study was to examine [Ca2+]i at
different temperatures in rainbow trout atrial myocytes. Previous studies have
shown that trout experience acute temperature fluctuations of as much as
±10°C either while transversing thermoclines or as a result of
diurnal changes in shallow streams (Reid
et al., 1997
; Matthews and
Berg, 1997
). Thus, we conducted our studies on 14°C-acclimated
rainbow trout at 7°C, 14°C and 21°C. By simultaneously quantifying
[Ca2+]i and ICa at each of these
temperatures, we were able to assess the effects of temperature on
[Ca2+]i and indirectly separate the effect of
temperature on extracellular and intracellular Ca2+ fluxes.
In fish hearts, ambient temperature also directly modulates heart rate
(Farrell and Jones, 1992). In
turn, changes in contraction frequency affect many of the cellular processes
that underlie the transient change in [Ca2+]i
(Møller-Nielsen and Gesser,
1992
; Hove-Madsen and Tort,
1998
). For example, increasing the contraction frequency from 0.2
Hz to 1.4 Hz causes significant reductions in the amplitude of ICa,
as well as reductions in both the amplitude and duration of the AP in trout
ventricular myocytes (Harwood et al.,
2000
). Because the frequency of stimulation will modulate cellular
Ca2+ cycling, it is critical to consider the physiologically
relevant contraction frequency associated with a particular temperature when
trying to understand the implications of temperature change on
[Ca2+]i in vivo. Thus, we examined
[Ca2+]i and ICa in trout atrial myocytes that
were stimulated with physiological APs at physiologically relevant contraction
frequencies for the applied temperature. We also examined the effect of
temperature on [Ca2+]i and ICa in response to
conventional square (SQ) voltage clamp pulses at slow (0.2 Hz) and
physiologically relevant frequencies to gain mechanistic insight and to
facilitate comparison of our results with earlier studies.
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Materials and methods |
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Isolated myocyte preparation
A detailed description of the rainbow trout myocyte preparation has been
published previously (Vornanen,
1998; Shiels et al.,
2000
). Briefly, myocytes were obtained by retrograde perfusion of
the heart, first with an isolating solution for 8-10 min and then with a
proteolytic enzyme solution for 20 min. After enzymatic treatment, the atrium
was removed from the ventricle, cut into small pieces and then triturated
through the opening of a Pasteur pipette. The isolated cells were stored for
up to 8 h in fresh isolating solution at 14°C.
Solutions
The isolating solution contained: NaCl, 100 mmol l-1; KCl, 10
mmol l-1; KH2PO4, 1.2 mmol l-1;
MgSO4, 4 mmol l-1; taurine, 50 mmol l-1;
glucose, 20 mmol l-1; and Hepes, 10 mmol l-1; adjusted
to pH 6.9 with KOH. For enzymatic digestion, collagenase (Type IA), trypsin
(Type III) and fatty-acid-free bovine serum albumin (BSA) were added to the
isolating solution. Standard internal and external solutions were used to
eliminate contaminating Na+ and K+ currents.
Tetrodotoxin (TTX, 1 µmol l-1) was included in all external
solutions to effectively eliminate fast Na+ currents
(Vornanen, 1998).
Cs+-based internal and external solutions with 15 mmol
l-1 tetraethylammonium chloride (TEA) in the pipette were used to
block K+ currents (Hove-Madsen
and Tort, 1998
). The extracellular solution for recording
ICa contained: NaCl, 150 mmol l-1; CsCl, 5.4 mmol
l-1; MgSO4, 1.2 mmol l-1;
NaH2PO4, 0.4 mmol l-1; CaCl2, 1.8
mmol l-1; glucose, 10 mmol l-1; and Hepes, 10 mmol
l-1; adjusted to pH 7.7 with CsOH. Additionally, 5 nmol
l-1 adrenaline was added to emulate the tonic level of adrenaline
circulating in the blood of resting rainbow trout in vivo
(Milligan et al., 1989
). The
pipette solution contained: CsCl, 130 mmol l-1; MgATP, 5 mmol
l-1; TEA, 15 mmol l-1; MgCl2, 1 mmol
l-1; oxaloacetate, 5 mmol l-1; Hepes, 10 mmol
l-1; EGTA, 0.025 mmol l-1; Na2GTP, 0.03 mmol
l-1; and K5-Fura-2, 0.1 mmol l-1; pH adjusted
to 7.2 with CsOH. All drugs, with the exception of TTX (Tocris, Bristol, UK),
were purchased from Sigma (St Louis, MO, USA).
Patch-clamp recordings
Whole-cell voltage-clamp experiments were performed using an Axopatch 1D
amplifier with a CV-4 1/100 headstage (Axon Instruments, Foster City, CA,
USA). Pipettes had a resistance of 2.2±0.30 M when filled with
pipette solution. Junction potentials were zeroed prior to seal formation.
Pipette capacitance (9.3±0.2 pF) was compensated after formation of a
G
seal. Mean series resistance was 6.5±0.3 M
. Membrane
capacitance (31.2±0.8 pF, N=76) was measured using the
calibrated capacity compensation circuit of the Axopatch amplifier. Signals
were low-pass filtered using the 4-pole lowpass Bessel filter on the
Axopatch-1D amplifier at a frequency of 2 kHz and were then analyzed off-line
using pClamp 6.0 software (Axon Instruments).
In a separate series of experiments, we recorded APs from rainbow trout
atrial myocytes (Fig. 1). Cells
were stimulated to elicit APs at 7°C, 14°C or 21°C and at a
frequency that corresponds to the resting heart rate of rainbow trout in
vivo at each of these temperatures. The temperature/frequency pairs were
as follows: 0.6 Hz at 7°C, 1.0 Hz at 14°C and 1.4 Hz at 21°C
(Tuurala et al., 1982;
Farrell et al., 1996
;
Aho and Vornanen, 2001
). APs
were elicited by the minimum voltage pulse able to trigger a self-sustained AP
(approximately 1 ms, 0.8 nA). The resultant AP waveforms were subsequently
used to provide a more physiologically relevant stimulus under which to
evaluate ICa and [Ca2+]i during temperature
change. In all experiments, the capacitive transients resulting from the AP
waveform were compensated for by the amplifier.
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Peak current amplitude of ICa was calculated as the difference between the peak inward current and the current recorded at the end of the depolarizing pulse. The charge (pC) carried by ICa was calculated from the time integral of the current and then normalized to cell capacitance (pC pF-1).
Fluorescent recordings
For each cell, after formation of the G seal, and before rupture of
the cell membrane, background fluorescence (approximately 40% and 25% of the
total signal at 340 nm and 380 nm, respectively) was recorded and then
subtracted from all subsequent measurements. Temperature change had no effect
on background fluorescence. Myocytes were alternately illuminated (optical
chopper, 0.2 kHz, 75 W Xenon arc lamp) at 340 nm and 380 nm with a high-speed
dual-wavelength filter-based illuminator (PowerFilter, Photon Technology
International, Brunswick, NJ, USA). Emitted light was filtered at 510 nm and
detected by a photon-counting photomultiplier tube, the output of which was
connected to a computer via an I/O board and saved to disk for later
analysis using the FeliX acquisition and analysis software (7-point smoothing
and trace averaging, Photon Technology International). The emission aperture
window was kept as small as possible and was focused close to, but out of view
of, the pipette tip to minimize any effect of cell movement during contraction
(cells typically contracted approximately 10-15% of the resting length).
The ratio of emitted fluorescence at 340 nm/380 nm (R) was
calculated to give an index of [Ca2+]i.
[Ca2+]i was calculated using the acquired fluorescent
ratios and calibration parameters derived both experimentally and through
calculation. To measure Rmax and Rmin,
cells were patched-clamped and perfused with an extracellular solution
containing metabolic inhibitors (rotenone, 2 µmol l-1; carbonyl
cyanide m-chlorophenyl-hydrazone (CCCP), 5 µmol l-1;
jodo acetic acid, 5 mmol l-1). Pipette and extracellular solutions
were the same as those described above except that nucleosides and sulphates
were omitted, respectively. To estimate Rmax, 10 µmol
l-1 Ca2+ ionophore (4-bromo A23187) and 1.8 mmol
l-1 CaCl2 were added to the perfusate. To estimate
Rmin, cells were then perfused with a solution containing
10 mmol l-1 EGTA. The values obtained are as follows:
Rmax=9.80±0.67,
Rmin=0.99±0.01, ß (signal fluorescence at 380
nm/background fluorescence at 380 nm)=4.62±0.44; N=17 (A.
Ryökkynen and M. Vornanen, unpublished data). The affinity constant
Kd of Fura-2 has been measured in vivo over a
range of temperatures (5-37°C) in enterocytes from the Atlantic cod
(Gadus morhua; Larsson et al.,
1999). We adjusted the Kd values obtained in
that study to match our experimental conditions of pH, buffer and ionic
strength by calculating the effects of pH, buffer and ionic strength on
1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA)
(Harrison and Bers, 1987
)
using the MaxChelator program (Bers et al.,
1994
). The calculated Kd values were as
follows: 395 nmol l-1 at 7°C, 366 nmol l-1 at
14°C and 336 nmol l-1 at 21°C. The fluorescence ratios
obtained during the experiments were converted into
[Ca2+]i using these Kd values and
the above measurements of Rmax, Rmin
and ß, as described by Grynkiewicz et al.
(1985
).
Experimental procedures
Atrial myocytes were superfused at a rate of 2 ml min-1 with
extracellular solution at 7°C, 14°C or 21°C. Each cell was tested
at one experimental temperature only. The extracellular solution was heated or
chilled by a water bath circuit before emptying into the recording chamber
(RC-26, Warner Instrument Corp., Hamden, CT, USA; volume 150 µl). A
thermocouple was placed inside the recording chamber and positioned no less
than 5 mm from the cell to ensure it was experiencing the desired temperature.
At the onset of each experiment, cells were stimulated with 25 physiological
APs at a frequency that corresponded to the in vivo heart rate of
trout at each temperature (see Fig.
1) to ensure that SR Ca2+ content was similar in all
cells (Shiels et al., 2002a).
However, the subsequent stimulation protocols may alter SR Ca2+
content as the experiment proceeds, which is a limitation of our approach.
Cells were first stimulated to contract with five SQ depolarizing pulses
(from -80 mV to +10 mV, 500 ms duration) at a frequency of 0.2 Hz
(SQ0.2Hz), and ICa and [Ca2+]i
were simultaneously recorded. The cells were then stimulated to contract with
five SQ depolarizing pulses at a frequency that corresponded to the in
vivo heart rate for rainbow trout at the given test temperature
(SQphysiol). SQ voltage-clamp protocols are vital for
characterizing channel kinetics and simplify interpretation of mechanistic
relationships, but they may have only limited relevance to the condition
in vivo (Linz and Meyer,
1998; Puglisi et al.,
1999
). Therefore, after stimulation under SQphysiol
conditions, the cells were stimulated with an AP waveform (see
Fig. 1) appropriate for the
test temperature and at a frequency appropriate for the test temperature
(APphysiol). ICa and [Ca2+]i were
simultaneously recorded throughout (see
Fig. 4). Although the
SQphysiol part of the protocol is of little physiological
significance, we believe that comparing the results obtained at physiological
stimulation frequencies from both a 500 ms SQ (SQphysiol) and a
physiological AP (APphysiol) allows us to evaluate the importance
of the AP and the possible limitations of the SQ pulse in assessing
physiological processes. The duration of each protocol was approximately 1 min
and we did not observe any changes in intracellular buffering by Fura-2 or
run-down of ICa over this time period.
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Statistical analysis
One-way repeated measures analysis of variance (RM ANOVA) were used to
compare the effects of changing the frequency and the stimulus waveform on
ICa and [Ca2+]i. One-way ANOVAs were used to
test the effects of temperature on ICa and
[Ca2+]i. Differences were considered significant at
P<0.05, as assessed by StudentNewmanKeuls (SNK)
post-hoc analysis. The mean values ± S.E.M. and the
statistical analysis for [Ca2+]i appear in the tables.
Five transients were recorded under each experimental pulse condition (i.e.
SQ0.2Hz, SQphysiol and APphysiol; see
Fig. 4). The diastolic
[Ca2+]i at the end of the 4th transient and the peak
[Ca2+]i of the 5th transient were used to represent a
steady-state value for each condition (Tables
1,2,3).
For future experiments, it would, however, be prudent to provide a longer
train of stimulation pulses, especially at warm temperatures, to ensure a
steady-state level had been attained. On the other hand, excursions by fish
across thermoclines can be very rapid and transitory in nature and therefore
ICa may not reach a steady-state in vivo before
temperature (and thus heart rate) change again.
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Results |
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During SQ pulse stimulation, we observed a temperature-dependent change in the amplitude of [Ca2+]i but no change in the charge density calculated by integrating ICa. This suggests that, in addition to Ca2+ flux across the SL, other Ca2+ flux pathways, most probably Ca2+ release from the SR, must be changing with respect to temperature in these cells. During AP stimulation, the reduced charge density of ICa at warm temperatures (Fig. 3B) may underlie the large percentage reduction in [Ca2+]i observed under these conditions (Fig. 2, Table 1).
Effects of stimulus change
The frequency and shape of the depolarizing stimulus profoundly altered
cellular Ca2+ dynamics. As a detailed example,
Fig. 4 illustrates the
frequency dependence and the influence of pulse shape on ICa and
[Ca2+]i for an experiment conducted at 21°C. During
SQ0.2Hz, diastolic Ca2+ concentrations returned to rest
between depolarizing pulses (Fig.
4A). When stimulation frequency was increased from
SQ0.2Hz to SQ1.4Hz, diastolic Ca2+
concentration increased significantly due to the long (500 ms) and
high-frequency (1.4 Hz) depolarization of the membrane (also see
Table 3). In addition,
[Ca2+]i increased by approximately 30%
(Fig. 4,
Table 1). The rate of rise also
increased significantly (from 0.73±0.11 µmol l-1
s-1 to 1.16±0.84 µmol l-1 s-1), and
the time required to reach the peak of the transient decreased during
SQ1.4Hz compared with that during SQ0.2Hz
(Table 2). When the stimulus
waveform was changed from SQ1.4Hz to AP1.4Hz,
[Ca2+]i and diastolic Ca2+ levels both
decreased significantly (Tables
1,
2), and the rate of rise of the
transient decreased significantly to a value (0.59±0.17 µmol
l-1 s-1) that was not significantly different from that
for the SQ0.2Hz pulse. Qualitatively similar effects of frequency
and stimulus shape were observed at 14°C and 7°C (Tables
1,2,3).
Thus, independent of temperature, an increase in stimulation frequency from
0.2 Hz to a physiologically relevant frequency for the experimental
temperature (SQphysiol) produced a large increase in both
[Ca2+]i and in diastolic Ca2+ levels with 500
ms SQ stimulation pulses. By contrast, the AP stimulus, which is much shorter
in duration, reduced [Ca2+]i and diastolic
Ca2+ levels compared with SQ pulses at the same frequency.
The amplitude of ICa was significantly smaller with an AP stimulus compared with a SQ stimulus at the same frequency (Figs 2,3,4; Table 1). The charge carried by ICa did not differ significantly at 14°C and 21°C but was greater at 7°C (Fig. 3B). This situation probably arose as a result of sustained Ca2+ influx during the AP plateau and because of Ca2+ influx occurring via the window current (see Fig. 2B and below).
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Discussion |
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Effects of temperature on [Ca2+]i and
ICa
With both SQ and AP pulses, [Ca2+]i was slowest and
largest at 7°C compared with at 14°C and 21°C. This observation
agrees with results from multicellular cardiac muscle studies from rainbow
trout, where isometric contraction is slower and stronger as temperature is
decreased (Driedzic and Gesser,
1994; Shiels and Farrell,
1997
; Aho and Vornanen,
2001
). These findings also agree with mammalian studies that
demonstrate an increase in the amplitude of the transient change in
[Ca2+]i with decreases in temperature
(Wang et al., 2000
). The small
amplitude of [Ca2+]i and the elevated diastolic
Ca2+ levels during physiological AP pulses at 21°C is striking
and is consistent with the finding that the maximum power output of trout
heart preparations in situ falls off at temperatures above 18°C
(Farrell et al., 1996
).
Indeed, the poor maximum cardiac performance at temperatures approaching the
upper incipient lethal temperature for salmonids (23-25°C;
Black, 1953
), coupled with the
observed reduction in maximum swimming capabilities of salmonids as
temperatures approach 21°C in both lab
(Brett, 1971
) and field
(Tierney, 2000
) studies, has
led to the suggestion (Farrell,
1997
) that cardiac failure may be a critical factor when salmon
are exposed to temperatures that exceed their preferred temperature
(approximately 15°C; Brett,
1971
). The implication of the present study is that the small
amplitude of [Ca2+]i in the cardiac myocyte may be
critical to setting the upper thermal regime in fishes. Thus, it follows that
data obtained from trout cardiac myocytes at room temperature should be
regarded with some caution.
Previous studies with trout atrial
(Shiels et al., 2000) and
ventricular (Kim et al., 2000
)
myocytes have shown a reduction in the peak current amplitude of
ICa with decreasing temperature, and our present study supports
these earlier findings. The similar Ca2+ influx via
ICa at 7°C, 14°C and 21°C probably resulted from a
combination of higher and faster peak ICa, offset by more rapid
ICa inactivation as temperatures increased (see
Fig. 2 and
Shiels et al., 2000
). A
similar lack of effect of temperature on the charge density of ICa
has been reported for rabbit myocytes at 25°C and 35°C
(Puglisi et al., 1999
). We
observed an increase in the outward current at the end of the stimulus pulse
at 21°C compared with at 14°C and 7°C (see
Fig. 2B), which is probably
attributable to increased Ca2+ influx via reverse-mode
sarcolemmal Na+/Ca2+ exchange (NCX;
Hove-Madsen et al., 2000
).
During AP stimulation, the charge density of ICa was reduced at
warm temperatures (Fig. 3B),
probably due to the shorter AP duration. This will have contributed to the
reduction in the transient change in [Ca2+]i observed
during stimulation at 21°C. However, certainly during the SQ pulse
stimulus protocol, and also during the AP stimulus protocol, the `mismatch'
between temperature-induced changes in SL Ca2+ influx and
[Ca2+]i suggests that temperature must modulate another
Ca2+ flux pathway in trout atrial myocytes.
Effects of increasing SQ pulse stimulation frequency on
[Ca2+]i
Increasing SQ pulse stimulation frequency (from SQ0.2Hz to
SQphysiol) caused an approximately 30% increase in
[Ca2+]i and an approximately 75% increase in the
diastolic Ca2+ level (Figs
2,
4; Tables
1,
3), indicating that more
Ca2+ enters the cytosol than exits via SR uptake or SL
efflux. The high-frequency application of long SQ pulses (500 ms) increased
Ca2+ influx via reverse-mode NCX, as evidenced by the
increased outward current at the end of the stimulation pulse (see
Fig. 4B, SQ1.4Hz).
In avian (Lee and Clusin,
1987) and mammalian cardiomyocytes
(Frampton et al., 1991
), the
frequency-dependent increase in [Ca2+]i has been
attributed, in part, to increased SR Ca2+ load and thus an
increased SR Ca2+ release. Increased Ca2+ influx
via NCX may have resulted in greater SR Ca2+ loading and
thus greater Ca2+-induced Ca2+ release, contributing
further to the increase in the amplitude of the transient change in
[Ca2+]i in the present study. An increase in the
frequency of field stimulation from 0.6 Hz to 1.0 Hz at 14°C increased
diastolic Ca2+ (approximately 25-50%) without changing the
amplitude of [Ca2+]i in trout ventricular myocytes
(Harwood et al., 2000
). In the
same study, the authors found that application of caffeine revealed a greater
SR Ca2+ content at 1.0 Hz compared with at 0.6 Hz, indicating that
SR Ca2+ content increased with pacing frequency but was not
releasable during a field-stimulated twitch
(Harwood et al., 2000
). The
differences between studies may indicate that a SQ pulse is a better trigger
for SR Ca2+ release than is field stimulation, where the membrane
is depolarized with an AP (see below). Indeed, Hove-Madsen et al.
(2001
) have demonstrated a
similar degree of CICR at 7°C and 21°C with SQ pulses in trout atrial
myocytes.
Effects of action potential stimulation on
[Ca2+]i
[Ca2+]i and diastolic Ca2+ levels
decreased dramatically when the SQ pulse was changed to an AP pulse at the
same frequency. This reflects a reduction in SL Ca2+ influx and
also possibly a reduction in CICR with AP pulses, as the smaller amplitude of
ICa during AP pulses (see Figs
2,
3) may be a less effective
trigger for CICR.
The myocytes were better able to manage diastolic Ca2+ levels at physiological frequencies with AP pulses (Table 3) but not as effectively as at 0.2 Hz. This suggests that the Ca2+ efflux mechanisms are not able to remove all of the Ca2+ effectively during diastole at physiological pacing frequencies. However, it is possible that resting diastolic Ca2+ concentration in the absence of stimulation and at 0.2 Hz is sub-physiological. The true `diastolic Ca2+ concentration' for fish hearts remains unknown and is worthy of future investigation. In any case, visual observations indicate that the cells were relaxing between stimulation pulses at physiological frequencies with AP pulses at all temperatures.
It has been estimated from trout ventricular myocytes at room temperature
that, when operating maximally (i.e. at or near Vmax), the
trout SR Ca2+-ATPase should be able to remove the total
intracellular Ca2+ transient between depolarizations at
physiological heart rates (Hove-Madsen et
al., 1998). However, our study reveals that, at room temperature,
the trout intracellular Ca2+ transient is small compared with
cooler temperatures and suggests that the SR Ca2+-ATPase is
unlikely to be operating near Vmax. Furthermore, estimates
of the maximal Ca2+ efflux rate of the rainbow trout NCX at room
temperature (Hove-Madsen and Tort,
2001
) suggest that the NCX can remove a physiological
Ca2+ transient from the cytosol in a few hundred milliseconds.
Based on those estimations, the NCX should have been able to remove all of the
cytosolic Ca2+ during the 700 ms between depolarizations at 1.4 Hz
in the present study at 21°C. The observed increase in the diastolic
Ca2+ levels in the present study are in contrast to these estimates
of Ca2+ efflux and may reflect exogenous buffering by Fura-2. The
in vivo buffering capacity of the fish myoplasm and its modulation by
temperature is unknown. However, because both contractility and the time
course of ICa were maintained in the presence of Fura-2, it is
unlikely that exogenous buffering was large relative to the measurements being
made. A comparison of the time to 50% relaxation of
[Ca2+]i with AP pulses in our study at 14°C
(approximately 250 ms) with the ratiometric results from the only other
published study of [Ca2+]i in fish hearts, which
examined field-stimulated trout ventricular myocytes loaded with Fura-2-AM at
15°C (approximately 230 ms; Harwood et
al., 2000
), suggests comparable time courses of the transients.
Additional studies are needed to resolve the physiological efficacy of trout
Ca2+ efflux pathways and the effect of exogenous buffering and
temperature change therein.
Effects of stimulus change on ICa
The frequency-dependent decrease in ICa at 21°C reported
here agrees well with previous studies that report a 15-20% decrease in peak
current amplitude over a similar frequency range at 21°C in trout myocytes
(Hove-Madsen and Tort, 1998;
Harwood et al., 2000
). These
responses are akin to the negative force-frequency response observed
independent of temperature in the myocardium of a number of fish species
(Driedzic and Gesser, 1985
;
Keen et al., 1994
;
Shiels and Farrell, 1997
; see
Shiels et al., 2002b
for a
recent review). The smaller amplitude of the peak current during an AP pulse
compared with during a SQ pulse is qualitatively similar to what we have found
previously in trout atrial myocytes
(Shiels et al., 2000
) and
similar to what has been described in mammals
(Linz and Meyer, 1998
;
Puglisi et al., 1999
).
The shape of the trout AP, in combination with the relatively depolarized
resting membrane potential (-50 mV), may have led to an overestimation of SL
Ca2+ flux via the ICa window current in the
present study (Fig. 2B). This
is because the late phase depolarization after the initial hyperpolarization
(Fig. 1) brings the membrane
potential close to the window current voltage range for ICa in
trout atrial myocytes (-40 mV to +30 mV, with a peak at -10 mV; see
Shiels et al., 2000). Recent
measurements in vivo suggest that the resting membrane potential in
rainbow trout atrial cells is approximately -65 mV
(Vornanen et al., 2002
). At
this membrane potential, the late phase depolarization would not enter the
window voltage range and thus ICa inactivation would be maintained.
The relatively depolarized membrane potential in isolated myocytes from our
study results from the loss of cholinergic tonus and also reflects the lower
density of inward rectifier K+ channel current (IK1) in
atrial compared with ventricular cells
(Vornanen et al., 2002
).
Future studies with isolated trout myocytes could consider including tonic
levels of acetylcholine when measuring APs to account for this.
Summary
Collectively, we have characterized some of the temperature-, frequency-
and waveform-dependent changes in [Ca2+]i and
ICa that exist in rainbow trout atrial myocytes. By calibrating the
Fura-2 signal, we were able to make meaningful comparisons of
[Ca2+]i at physiologically relevant temperatures for
rainbow trout. We have shown that increased temperature causes a decrease in
[Ca2+]i. This is the first cellular observation of a
temperature-dependent change in [Ca2+]i in fish myocytes
and may help to explain the well-known negative inotropic effect of warm
temperatures on trout heart function. Our results suggest that the
temperature-dependent changes in [Ca2+]i are not solely
dependent on temperature-dependent changes in ICa. Therefore,
temperature and frequency modulation of SR Ca2+ accumulation and
release may be involved, and future studies should be directed at
understanding these mechanisms. Finally, we have shown that the amplitude of
[Ca2+]i and the level of the diastolic Ca2+
concentration is dependent on the shape and the rate of stimulation. At all
temperatures, cells were better able to maintain diastolic Ca2+
levels at physiological frequencies with AP pulses compared with 500 ms SQ
pulses. This suggests that temperature-dependent modulation of the shape of
the AP is important for cellular Ca2+ regulation during temperature
and frequency changes in rainbow trout heart.
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Acknowledgments |
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References |
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