Vol. 282, Issue 4, R1200-R1209, April 2002
Electrophysiological properties of rainbow trout cardiac
myocytes in serum-free primary culture
Antti
Nurmi and
Matti
Vornanen
University of Joensuu, Department of Biology, 80101 Joensuu, Finland
 |
ABSTRACT |
A
low-density primary culture of trout ventricular myocytes in serum-free
growth medium was established and maintained for up to 10 days at
17°C. The myocytes retained their normal rod shaped morphology,
capacitive surface area of the sarcolemma (SL), and contractile
quiescence. However, sarcolemmal cation currents changed significantly,
some permanently, some transiently, after 8-10 days of culture.
TTX-sensitive sodium current (INa) and
Ba2+-sensitive background inward rectifier potassium
current (IK1) were permanently depressed to
24-28% of their control density measured in freshly isolated
myocytes. In contrast, L-type calcium current
(ICa) was only transiently downregulated; after
2-3 days in culture, the density of the current was 32% of the
control and recovered to the control value after 8-10 days in
culture. The changes in membrane currents were reflected in the shape
of the action potential (AP). After 2-3 days in culture, maximal overshoot potential and resting potential were significantly reduced, and the durations of the AP at 50 and 90% repolarization were significantly increased. These changes became significantly more pronounced after 8-10 days of culture, with the exception of AP duration at 50% repolarization level. The shortening of the early plateau phase may reflect an additional change to an outward current, presumably the rapid component of the delayed rectifier
(IKr). Although the present findings indicate
that fish cardiac myocytes can be maintained in serum-free primary
culture for at least 10 days at 17°C, some but not all of the
electrophysiological characteristics of the myocytes change markedly
during culture. The changes in ion currents were not due to loss of
sarcolemmal membrane and therefore are likely to represent altered
expression of cation currents as an adaptive response to culture conditions.
cultured cardiac cells; electrophysiology; fish heart; cation
currents; action potential
| |
INTRODUCTION |
CELL CULTURE OFFERS A
SIMPLE environment compared with animal body and therefore a
potentially useful in vitro system to examine adaptation mechanisms of
the cardiac myocyte to altered environmental conditions. Unlike the
complexity of whole animal models, cell culture allows better control
of external factors that could help in revealing cause-effect
relationships of physiological adaptation processes. Indeed,
ventricular myocytes of adult mammals can be maintained in
serum-supported culture media for weeks to months (for review, see Refs
13, 16). However, for studying adaptational changes, the presence of
serum, which contains a number of growth factors, hormones, and other
undetermined components, makes the culture system a nondefined
environment and therefore limits its value as a research tool. In
contrast, a serum-free cell culture provides a better environment to
study the adaption of cardiac myocytes to changing conditions. The goal
here was to ascertain, for the first time, whether 1) fish
cardiac myocytes could be held in serum-free culture and 2)
to examine the electrophysiological stability of fish myocytes held in
serum-free conditions.
Mammalian cardiac myocytes have been recently cultured for up to 6 days
in serum-free media (15). The adult mammalian cardiac myocytes show several morphological, contractile, and
electrophysiological changes during the culture and some of the
electrophysiological changes seem to be closely associated with
ultrastructural transformations induced by culture conditions
(16). For example, the extensive t-tubular system of the
adult cardiac myocytes is practically lost during cell culture,
resulting in a marked reduction of sarcolemmal surface area (5,
15). The loss of t-tubules may also be responsible for the
suppression of inward rectifier K+ current
(IK1) in mammalian cell cultures, as the
channels are almost exclusively located in the t-tubular membrane
(5, 15, 22). However, fish cardiac myocytes lack t-tubules
(18), and therefore the prediction is that the
electrophysiological modifications during culture will be smaller. To
assess preservation of ion channel currents, action potential (AP)
shape and capacitive cell surface area, ventricular myocytes of the
rainbow trout heart were cultured in serum-free medium for up to 10 days.
Although there were no changes in capacitive cell surface area during
culture, a sequence of changes in sarcolemmal cation currents occurred,
suggesting that culture-induced modifications of ion currents are not
due to the loss of sarcolemmal membrane area, but rather due to the
altered expression of channel proteins under culture stresses.
| |
MATERIALS AND METHODS |
Isolation of ventricular myocytes.
Female rainbow trout (n = 13, body mass between 101 and
154 g) were obtained from a local fish farm (Kontiolahti) and
acclimated to 17-18°C for at least 3 wk before the experiments.
Fish were maintained under a constant 15:9-h light-dark photoperiod in
500-liter stainless steel tanks provided with circulating (~0.5
l/min) and aerated tap water. The fish were fed daily with a commercial
trout chow (Ewos; Turku, Finland). Ventricular myocytes were obtained by enzymatic isolation procedure described in detail in earlier publications (23, 24). Isolation procedures were conducted with aseptic instruments and equipment to avoid microbial
contamination. All solutions were sterilized and filtered before use.
Briefly, the fish was stunned with a blow to the head and the spine was cut. The heart was quickly dissected and perfused retrogradely with
oxygenated (100% O2) nominally Ca2+-free
isolation solution (in mM: 100 NaCl, 10 KCl, 1.2 KH2PO4, 4 MgSO4, 50 taurine, 20 glucose, and 10 HEPES at pH 6.9) for 8-10 min followed by
enzymatic digestion with collagenase IA (Sigma, 0.75 mg/ml) and trypsin
III (Sigma, 0.5 mg/ml) for 20 min at room temperature (~20°C). The
isolation solution was supplemented with fatty acid-free BSA (Sigma;
0.75 mg/ml) and antibiotics (50 IU/ml of streptomycin and penicillin).
The enzymatically digested ventricle was minced with scissors in small
pieces, and single myocytes were liberated in fresh and oxygenated
isolation solution by triturating ventricular pieces through the
opening of a Pasteur pipette. The number of viable cells, determined by
Trypan blue (Sigma-Aldrich) exclusion in a hemocytometer
(Bürker, Kebolab), accounted for 50-70% of the whole cell
population. Isolated cells were suspended in sterile isolation solution
at the density of ~50,000 cells/ml.
Cell culture.
The so-called "rapid attachment" method (13) was used
to establish a low-density primary culture of ventricular cells. Trout ventricular myocytes were grown on clean, uncoated, round glass coverslips (12 mm in diameter) in plastic culture dishes
(Cellstar7; 35/10 mm; Greiner Labortechnik). Four
coverslips with a total number of ~5,000 myocytes were placed in each
dish and covered to prevent drying out. Thirty minutes later, when the
cells were firmly attached on glass, 1.8 ml culture medium (DMEM/F-12
with L-glutamate and 15 mM HEPES; Sigma) was added in each
dish. Culture medium was supplemented with 12 mM NaHCO3,
antibiotics (streptomycin and penicillin 100 U/ml), 30 mM ARA-C
(Sigma), and adjusted with 0.5 N NaOH to pH 7.6. Culture dishes were
placed in a temperature-controlled (17°C) incubator (Calaxy, RS
Biotech) with regulated CO2 (2%) and O2 levels
(19%). Trout ventricular myocytes were grown for a maximum of 10 days,
and the medium was changed every 3 days during the culture. The
electrophysiological experiments were conducted with cells cultured for
2-3 and 8-10 days and compared with those performed on
freshly isolated myocytes (control).
Electrophysiological measurements.
Standard patch-clamp methods in whole cell configuration
(10) were used to record cation currents and APs as
reported in detail elsewhere (23, 24, 26). Before the
patch-clamp experiments began, the cells were washed free of the
culture medium by immersing the glass coverslips in
K+-based Tyrode solution (see below) for 1-2 h at room
temperature (~20°C). The coverslip with attached cells was placed
in the recording chamber (RC-26, Warner Instrument), and the cells were
observed with an inverted microscope (Nikon Eclipse 200). Cells were
superfused (~2 ml/min) with an external solution of appropriate
composition for the parameter under study. For the recording of sodium
current (INa) and L-type calcium current
(ICa-L), a Cs+-based external
solution (in mM: 150 NaCl, 5.4 CsCl, 1.8 CaCl2, 1.2 MgCl2, 10 HEPES, and 10 glucose, pH 7.6 adjusted with NaOH) was used. Nifedipine (10 µmol/l; Sigma) and TTX (1 µmol/l; Alomone Labs, Israel) were included, respectively, when recording
INa and ICa-L. APs and
K+ currents were recorded in K+-based external
solution (in mM: 150 NaCl, 5.4 KCl, 1.8 CaCl2, 1.2 MgCl2, 10 HEPES, and 10 glucose, pH 7.6 adjusted with KOH). Nifedipine (10 µmol/l; Sigma) and TTX (1 µmol/l) were added to this
solution when K+ currents were examined. APs were
stimulated at 0.2 Hz with 1-ms current pulses just exceeding the AP
threshold (~1 nA).
The pipette solution for INa and
ICa-L recordings was as follows (in mM): 130 CsCl, 15 TEA, 10 HEPES, 5 MgATP, 1 MgCl2, 5 oxaloacetate, 5 EGTA, pH 7.2 adjusted with CsOH. For the recording of K+
currents, the pipette solution contained (in mM) 140 KCl, 4 MgATP, 1 MgCl2, 5 EGTA, 10 HEPES, pH 7.2 adjusted with KOH. Current
clamp recording of APs was performed with a pipette solution that
contained (in mM) 140 KCl, 5 Na2ATP, 1 MgCl2,
10 HEPES at pH 7.2 adjusted with KOH.
Glass pipettes were pulled from borosilicate capillaries (Modulohm)
using a two-stage vertical puller (Narishige MF 83). Pipette resistance
was 2-3 M
when filled with internal solutions. In AP
recordings, the potential difference (~7 mV) between the bath solution and the pipette solution was corrected in the final results. Voltage and current clamp experiments were made using an Axopatch 1D
amplifier (Axon Instruments) equipped with a CV-4 1/100 head stage
(Axon Instruments). Junction potentials were zeroed before formation of
the gigaohm seal. The pipette capacitance (4-6 pF) was compensated
for after the seal formation. The patch was ruptured by delivering a
short (0.1-1 ms) voltage pulse (1.5 V) to the cell. Capacitive
transients were eliminated by iteratively adjusting the calibrated
series resistance and whole cell capacitance circuits. The cell
capacitance was read directly from the dial of the amplifier. APs and
INa recordings were low-pass filtered at 10 kHz,
inward rectifier potassium current (IK1) and
ICa-L at 2 kHz. Current traces were sampled with
an analog-to-digital converter (TL-1 DMA, Axon Instruments) and stored
on the hard drive of a personal computer for offline analysis using the
pClamp 6.04 software (Axon Instruments).
Data analysis.
Kinetic parameters of steady-state activation as well as
current-voltage relationships for both INa and
ICa-L were obtained from square pulse recordings
(see RESULTS). Activation voltage-dependence, d
(V), was determined as
normalized Na+ or Ca2+ conductance,
d(V) = gNa or
Ca-L/gmax, where gmax is
the maximum value of INa or
ICa-L conductance. The voltage dependence of the
peak conductance for both currents was calculated using the equation
|
(1)
|
where gNa or Ca-L is Na+ or
Ca2+ conductance, INa or Ca-L is the
peak Na+ or Ca2+ current at a given potential
(V), and Vrev is the apparent reversal potential
obtained by extrapolating the ascending portion of the current-voltage
(I-V) relationship of Na+ or Ca2+
current to zero current. Steady-state activation parameters were obtained by fitting the obtained data to Boltzmann equation
![d<SUB>∞</SUB>(V)=1/{1+exp[<IT>V</IT><SUB>0.5</SUB><IT>−V</IT>)<IT>/k</IT>]}](/content/vol282/issue4/fulltext/R1200/img002.gif)
|
(2)
|
where V0.5 is the half-activation
potential and k is the slope factor.
Statistical analysis.
Statistical comparisons between freshly isolated trout ventricular
cells (control) and from both groups of cultured cells were made by
using a one-way ANOVA followed by Dunnet's post hoc analysis. In the
case of unequal variances between groups, a two-tailed Student's
t-test was used. A P value of 
0.05 was
considered as statistically significant.
| |
RESULTS |
Cell morphology during culture.
Trout ventricular cells adhered firmly on clean, uncoated glass
coverslips. After 1-2 h of plating, the myocytes were so firmly attached to the glass that removal of an individual cell was impossible without breaking it. The majority of the ventricular cells maintained their rod shape morphology for 8-10 days of culture, even though the cell endings became slightly rounded in some cells (Fig.
1). Extensive branching or spreading of
the cells on the glass was rare. No cell-to-cell contacts were
observed, and the myocytes did not beat spontaneously.
View larger version (77K):
[in this window]
[in a new window]
|
Fig. 1.
Photomicrographs of freshly isolated (A),
3-day cultured (B), and 10-day cultured (C)
cardiac myocytes of the rainbow trout ventricle. The cells were fixed
in Bouin and stained with hematoxylin-eosin. The scale bar is 50 µm.
|
|
Culture conditions did not significantly affect sarcolemmal surface
area as whole cell capacitance was unchanged during the 10-day culture
period (Fig. 2).
View larger version (13K):
[in this window]
[in a new window]
|
Fig. 2.
A box plot of whole cell capacitance in freshly isolated
and cultured cardiac myocytes of the rainbow trout ventricle. There
were no statistical differences between the 3 groups of cells
(P > 0.05) (Kruskall-Wallis test).
|
|
Changes in resting membrane potential and APs during culture.
APs recorded from freshly isolated cells displayed a rapid upstroke, a
large overshoot (OS) with a maximum value of +59.9 ± 3.79 mV
(n = 15) and a long plateau phase between +40 to +10 mV
before repolarization (Fig. 3). The
duration of the AP (APD) at 50 and 90% repolarization was 262.6 ± 21.2 and 306.6 ± 28.7 ms (n = 15),
respectively. The mean resting membrane potential (RP) of the
freshly isolated myocytes was 
75.2 ± 0.82 mV. After 2-3
days of culture, changes in RPs and APs were evident in trout ventricular myocytes (Fig. 3). RP was less negative, OS was reduced, and APD was markedly increased. Subsequent recordings from cells cultured for 8-10 days (Fig. 3) revealed progressively more
depolarized RPs, decreased OSs, and increased APD at 90%
repolarization level. There was, however, no statistical difference in
APD at 50% repolarization level between freshly isolated cells and
cells cultured for 8-10 days (Table
1).
View larger version (11K):
[in this window]
[in a new window]
|
Fig. 3.
Examples of action potentials from freshly isolated
(control) and cultured cardiac cells of the rainbow trout ventricle.
Note the greater variability of action potential (AP) shapes in
cultured cardiac myocytes.
|
|
View this table:
[in this window]
[in a new window]
|
Table 1.
Characteristics of action potentials in freshly isolated and cultured
ventricular myocytes of the rainbow trout heart
|
|
Changes in INa during culture.
In freshly isolated myocytes, 10-ms depolarizing pulses from a holding
potential (HP) of 80 mV to various potentials between 120 and +60
mV generated a fast, transient, and large (6-10 nA) inward current
(Fig. 4). This current was completely
blocked by a relatively low concentration (1 µmol/l) of TTX and,
therefore, represent typical INa of fish cardiac
myocytes (23). The I-V relationship was bell
shaped, with a threshold around 50 mV, the peak current density at
10 mV, and the apparent reversal potential at +90 mV (Fig. 4; Table
2). The potential for half-maximal activation (V0.5) occurred at 25.97± 1.24 mV
with a slope (k) of 5.38 ± 0.15 (n = 5). The inactivation rate of current at peak INa
(10 mV) was fitted to a double exponential function, which gave mean
time constants of 0.177 ± 0.01 ms and 0.867 ± 0.01 ms for
the fast (
f) and slow component (s) of
current decay, respectively (Fig. 5).
View larger version (27K):
[in this window]
[in a new window]
|
Fig. 4.
Effect of serum free primary culture on sodium current
(INa) of trout ventricular cells. A:
families of INa in the absence (left)
and presence (right) of 1 µM TTX at different membrane
potentials. Note the different y-axis scaling for each group
and the different x-axis scaling between freshly isolated
and cultured cardiac cells. B: current-voltage relationship
(left) of INa density and
steady-state activation (right) of
INa for freshly isolated myocytes (control), 2- to 3-days cultured cells (2-3 d), and 8- to 10-days cultured cells
(8-10 d). The results are means ± SE of 5 cells for all
groups.
|
|
View larger version (24K):
[in this window]
[in a new window]
|
Fig. 5.
Kinetics of INa inactivation.
A: original recordings of peak INa
and double exponential fits to the recorded currents from freshly
isolated and cultured myocytes. Mean ± SE results for fast
(B) and slow (C) time constants of inactivation.
The results are means ± SE of 5 cells for all groups.
* Statistically significant difference from the value of freshly
isolated cells.
|
|
In myocytes cultured for 2-3 days, the density of
INa was only 53% of that measured in freshly
isolated cells, and the peak current density was shifted to more
positive voltages (Fig. 4; Table 2). The slope factor of the
steady-state activation curve was also increased, but the
V0.5 was not significantly different from the
value of freshly isolated cells. Although the time constants of current
inactivation at the peak density of INa (0 mV)
were somewhat slower than in control cells, the differences were not statistically significant at this phase of culture (Table 2).
After 8-10 days in culture, the peak density of
INa was further shifted to +30 mV, and the
current density was further reduced to 24% of the value in freshly
isolated myocytes (Fig. 4). The voltage-dependence of steady-state
activation (V0.5) was now significantly shifted
to the right, and the slope remained less steep than in freshly
isolated myocytes (Fig. 4). The rate of current inactivation in
myocytes cultured for 8-10 days was also much slower than in freshly isolated cells (Table 2).
The progressive decrease in density of INa
correlated strongly with the diminishing OS of the AP during the
culture (Fig. 6).
View larger version (19K):
[in this window]
[in a new window]
|
Fig. 6.
Correlation between maximum overshoot potential of the
action potential and the peak density of INa.
The results are means ± SE of 5 cells for each group.
|
|
Changes in IK1 during culture.
K+ currents were elicited in the presence of TTX and
nifedipine by square-wave voltage steps (500 ms) from the HP of 80 mV to various potentials between 120 and +20 mV. Hyperpolarizing voltage
steps between 20 and 80 mV elicited a noninactivating, Ba2+-sensitive inward current with nearly linear
I-V relationship (Fig. 7). The
slope conductance of the current in freshly isolated myocytes was
0.626 ± 0.046 nS/pF. Extrapolation of this linear current to zero
current level gave a reversal potential of about 73 mV. At the
positive side of the reversal potential, the current exhibited strong
inward rectification. On the basis of the inward rectification and
sensitivity to Ba2+, this current represents the background
inward rectifier (IK1).
View larger version (26K):
[in this window]
[in a new window]
|
Fig. 7.
Effect of serum-free culture on K+ currents
of rainbow trout ventricular myocytes. Representative recordings from
freshly isolated cells (A), 2- to 3-day cultured cells
(B), and 8- to 10-day cultured cells (C) in the
absence (left) and presence (right) of 0.5 mM
BaCl2. Note that in 8- to 10-day cultured myocytes there is
a Ba2+-resistant current component. Mean results (±SE)
from 8-10 cells are shown in D. The currents were
triggered from a holding potential of 80 mV for 500 ms to various
potentials between 120 and +20 mV. The currents were measured at the
end of the 500-ms pulse. Note the different y-axis scaling
in A, B, and C.
|
|
Subsequent recordings in cultured myocytes revealed a dramatic
reduction of the IK1 density (Fig. 7). Compared
with freshly isolated cells, after 2-3 days in culture the slope
conductance was <36% (0.228 ± 0.056 pS/pF; P < 0.05, n = 9) and after 8-10 days in culture only
28% (0.175 ± 0.066 pS/pF; P < 0.05, n = 10) of the value in freshly isolated cells. The
changes in slope conductance of IK1 and RP were
strongly correlated (Fig. 8).
View larger version (19K):
[in this window]
[in a new window]
|
Fig. 8.
Correlation between resting potential and sarcolemmal
K+ conductance. The results are means ± SE of
8-10 myocytes.
|
|
There was a small but significant increase in outward K+
current density between 0 and +20 mV in myocytes cultured for 8-10 days compared with freshly isolated ventricular myocytes. This was
probably due to a time-dependent outward current that had developed
after 8-10 days in culture (Fig. 9).
The time-dependent outward current was only slightly reduced by 0.5 mM
Ba2+ and it was not blocked by either Cd2+ (25 µmol/l) or verapamil (10 µmol/l) (data not shown).
View larger version (15K):
[in this window]
[in a new window]
|
Fig. 9.
A: left part of Fig. 7D
expanded to indicate the presence of the outward K+ current
in 8- to 10-day cultured myocytes. * Significantly different
(P < 0.05) from the value of freshly isolated cells.
B: an example of a current activated by depolarization from
80 mV to +20 mV in a 8- to 10-day cultured myocyte.
|
|
Changes in ICa-L during culture.
Ca2+ currents were recorded by clamping the membrane
potential for 500 ms from the HP of 50 mV to various potentials
between 50 and +60 mV. In freshly isolated trout ventricular
myocytes, a small inward current with inverted bell-shape
I-V relationship was recorded. The current activated at
around 40 mV, attained peak density at +10 mV, and finally approached
zero current level at +60 mV (Fig. 10).
The current was blocked by 10 µmol/l nifedipine, 25 µmol/l
Cd2+, and 10 µmol/l verapamil (data not shown). Analysis
of steady-state activation parameters gave V0.5
of 6.63 ± 0.53 and k of 6.22 ± 0.07 (n = 5). The rate of current inactivation was
biexponential with fast and slow time constants of 27.7 ± 2.3 and 95.6 ± 7.3 ms, respectively, n = 8)
(Fig. 11).
View larger version (25K):
[in this window]
[in a new window]
|
Fig. 10.
Effect of serum-free primary culture on L-type
Ca2+ current. A: representative recordings of
currents and their block by 10 µM nifedipine in freshly isolated and
cultured ventricular myocytes. Current-voltage relationship
(B) and steady-state activation (C) of
ICa of the 3 cell groups. The results are
means ± SE of 5-19 cells.
|
|
View larger version (29K):
[in this window]
[in a new window]
|
Fig. 11.
Kinetics of ICa inactivation at
different phases of serum-free culture. A: representative
recordings of ICa (dots) and their double
exponential fits (continuous lines). Mean ± SE results for the
fast (B) and slow (C) time constants of current
decay, respectively. * Significantly different (P < 0.05) from the value of freshly isolated cells.
|
|
During the initial 2-3 days of serum-free culture, the peak
density of ICa-L was reduced by 36%, and the
I-V relationship was shifted to the right. Voltage parameter
of steady-state activation was also shifted to more positive
potentials, but the slope factor was not changed (Table
3). The fast component of current
inactivation (f) was unchanged during the first 2-3
days in culture, but the slow component (s) increased
significantly during this period (Table 3).
After 8-10 days of culture, the density of
ICa-L had recovered to the level of freshly
isolated myocytes. However, the steady-state activation remained
shifted to the right, the slope had also increased significantly, and
inactivation rate constants (f and s)
were increased significantly compared with the freshly isolated myocytes.
| |
DISCUSSION |
Myocyte culture offers a homogenous population of single cells
that can be examined in a controlled environment. In principle, the
experimenter can change a single factor at a time and record its
influence on electrical activity of the myocyte. This requires that the
cells are grown in a defined culture medium in a constant physical
environment and that the electrophysiological changes associated with
myocyte adaptation to culture environment are known. In an ideal case,
after an adaptation phase the electrophysiological properties of the
cell will achieve a steady state, which can be used as a control for
the experimental manipulation.
Two basic methods have been used previously for culturing mammalian
cardiac myocytes (see Refs 13 and 16). In the redifferentiation method,
the myocytes lose their normal rod shaped morphology after a short time
in culture and begin to contract spontaneously in the absence of
external stimulation. First, the cells quickly dedifferentiate to a
fetal phenotype and then begin to differentiate back toward an adult
phenotype. Spontaneous activity and altered morphology indicate that
myocytes have lost their native in vivo properties. In contrast, in the
rapid-attachment method the cells start to grow in the adult
differentiated state and therefore retain their normal gross morphology
and contractile quiescence for a longer time before starting a gradual
transformation (dedifferentiation). Another major advantage of the
rapid-attachment method is that cells can be grown in defined culture
medium without serum.
The present experiments indicate that trout ventricular myocytes,
prepared with the rapid-attachment method, can be maintained in a
serum-free primary culture for at least 10 days. The cells essentially
retained their spindle shape in vivo morphology and did not show any
visible contractions or spontaneous electrical activity. Thus, in
regard to gross morphology and contractile and electrical quiescence,
the cultured cardiac myocytes of the trout heart in serum-free medium
seem to retain their native properties and therefore closely resemble
cultured cardiac cells of the mammalian heart prepared with the
rapid-attachment method (13).
Mammalian cardiac myocytes, however, reorganize their sarcolemma and
undergo a selective and extensive loss of t-tubular membrane resulting
in marked reduction in whole cell capacitance, i.e., sarcolemmal
surface area (5, 14, 15). We predicted and found that
there were no changes in whole cell capacitance in trout ventricular
myocytes during the 10-day culture, probably due to the absence of
t-tubules in fish cardiac myocytes. Nevertheless, this did not prevent
prominent changes in the electrophysiological properties of the trout
myocytes during culture.
Changes in RPs and APs during culture.
AP reflects the function of all voltage- and time-dependent ion
currents of the cardiac myocyte and is therefore a sensitive indicator
for any changes that might occur in underlying membrane currents during
the culture period. Freshly isolated ventricular myocytes of the
rainbow trout exhibited stable RPs and robust APs with similar
overshoot, amplitude, and duration as recorded by others in the same
species under similar experimental conditions (11).
However, during the serum-free primary culture, RPs depolarized, OS and
amplitude of AP decreased, and the duration of AP significantly increased. These changes in membrane potential of the cultured trout
ventricular myocytes are surprisingly similar to the changes found in
mammalian cardiac myocytes under culture conditions and over the same
time period (8, 15, 17, 19). For example, the depression
of the early plateau occurs in rabbit ventricular myocytes after 6 days
of culture in serum-free medium (15). Interestingly,
feline and rabbit ventricular myocytes cultured for 2 to 14 days in
serum-supplemented media exhibit early after depolarizations (EADs)
(19, 23). In serum-free media, EADs were not observed in
trout ventricular myocytes (present study) or in mammalian ventricular
cells (6, 15), suggesting the possibility that EADs are
due to serum-dependent changes in the expression or function of ion
channels. Regardless, the properties of cells grown in serum-free media
are closer to the native in vivo state of cardiac myocytes.
Changes in IK1 during culture.
Inward rectifier K+ channels maintain a stable RP and
contribute to the fast final phase of AP repolarization in cardiac
myocytes by allowing K+ efflux across the SL in the form of
background potassium current, IK1.
The high density of IK1 in freshly isolated
cells is consistent with our own observations in ventricular myocytes
of the rainbow trout acclimated to 17°C (26). The
background inward rectifier K+ current is severely and
quickly depressed during serum-free culture of the trout myocytes,
which is a routine finding in mammalian cardiac cell culture
preparations (3, 5, 15, 19, 23). The depression in the
density of IK1 in cultured mammalian cardiac myocyte is the result of a reduction in the number of active channels (23). On the basis of the delay of block and unblock of
IK1 by extracellular Ba2+ applied
rapidly to the cells, it is suggested that the inward rectifier
K+ channels of the mammalian ventricular myocytes are
located mainly in t-tubules (5). Thus the loss of a
greater part of the t-tubules during primary culture could explain the
culture-induced depression of IK1 in mammalian
myocytes (14, 15). The same conclusion cannot, however, be
made for the depression of IK1 in trout
ventricular myocytes that are devoid of t-tubules (18). It
is possible that some dedifferentiation toward the phenotype of
embryonic myocytes occurred during culture of trout ventricular
myocytes. Dedifferentiation could in principle cause the depression of
IK1 as the expression of the inward rectifier
channel is developmentally regulated and less well developed in early
developmental stages (3, 28, 29). It is also possible that
culture conditions decrease the rate of protein synthesis and/or
increase the rate of protein degradation with subsequent depression in
the number of functional channels.
There was also a significant increase in outward K+ current
at positive voltages (from 0 to +20 mV) in trout ventricular myocytes cultured for 8 to 10 days. The increase in outward current is accounted
for by the time-dependent and Ba2+-resistant current that
appears in later phases of culture. The time and voltage dependence of
this current when superimposed on the background inward rectifier
IK1 gave a superficial impression of a
Ca2+ current and therefore we assumed that it might be a
nifedipine-insensitive Ca2+ current. The resistance of this
current to inhibition by verapamil and Cd2+ suggests that
it is not carried by Ca2+ channels. Voltage dependence and
time course of activation of this current were similar to those of the
rapid component of the delayed rectifier K+ current
(IKr), which we have characterized in trout
atrial and ventricular myocytes (26). The density of this
current is small in ventricular myocytes of trout acclimated to 17°C
but it is upregulated when the fish are exposed for longer periods to
the cold (4°C) (25). Therefore, it is possible that in
serum-free culture the ventricular myocytes undergo a similar
physiological change as the ventricular myocytes of the rainbow trout
during cold acclimation.
It seems plausible that the changes in the expression of these two
functionally similar K+ currents were coupled so that
depression of one current results in upregulation of another, thereby
maintaining a balance between repolarizing and depolarizing currents in
cardiac myocytes. The two currents are not, however, functionally
identical, and the mutual replacement will probably alter excitability
of the myocytes. Clearly, more studies are needed to resolve the
physiological consequences of this altered balance in repolarizing
K+ currents in trout ventricular myocytes and the factors
that regulate their expression.
Changes in INa.
TTX-sensitive INa causes the fast upstroke and
prominent OS of the cardiac AP. The properties of
INa in trout ventricular myocytes are in some
respects different from those of mammalian cardiac myocytes. The
Na+ channels of the fish heart are more TTX sensitive than
their mammalian counterparts as 1 µM TTX completely blocks
INa in fish cardiac myocytes (24)
but not in mammalian cardiac cells. Voltage dependence of
INa is also different; the threshold voltage of INa activation and V0.5
of steady-state activation are more positive in trout ventricular
myocytes than in mammalian cardiac myocytes.
In trout ventricular myocytes, INa was
dramatically decreased during culture. The correlation between the
density of the INa and the OS of AP suggests
that progressive depression of OS is due to the diminished
INa. In addition to the decreased
INa, the low RP of cultured cardiac cells may
also diminish the OS, because depolarization reduces the number of
Na+ channels that are available for opening.
In mammals, the cardiac Na+ channels seem to be sensitive
to culture environment, although the response depends on the culture conditions (20). In trout myocytes,
INa was severely depressed even though the
capacitive area of the SL stayed constant for the whole culture period.
Therefore, the reduction in INa must be due to
changes in the number of functional Na+ channels or in the
conductance of individual channels. Similar for K+channels,
the number of Na+ channels might be reduced due to the
depression of protein synthesis or increase in the rate of degradation
of the existing channels.
In addition to the depression of INa density,
there were also very prominent changes in the kinetics of
INa. I-V relationship was shifted to
the right, and the activation and inactivation kinetics became slower
when the cells were grown in culture medium. On the extracellular side
of the membrane, Na+ channel proteins are conjugated with
carbohydrate moieties, which may have a role in ion channel function
(see Ref. 4). Sialic acids, which form terminal components
of oligosaccharide chains of glycolipids and glycoproteins, are
considered to be especially important (1, 21). Indeed, a
decrease in the amount of sialic acid on the Na+ channel
shifts all voltage-dependent gating parameters such that channels
require larger depolarizations to gate (2). Enzymes used
for myocyte isolation may also shift the voltage dependence of
INa (7). Therefore, we cannot
exclude the possibility that our method of myocyte isolation or culture
conditions might have affected sarcolemmal glycolipids and
glycoproteins and caused the positive shift in
INa voltage dependence during culture, e.g., by
changing the surface charge.
Activation and inactivation rate of INa
decreased during the whole culture period in rainbow trout ventricular
myocytes. An increase in the rate of activation and inactivation
kinetics of INa has been found in cultured
ventricular myocytes of the cat heart (20), whereas no
changes were evident in the kinetics of INa in
human atrial myocytes during a 5-day culture period (9).
In regard to the kinetics of INa, cultured trout
myocytes clearly differ from mammalian cardiac myocytes. This might be due to the fact that fish cardiac myocytes express a different (TTX
sensitive) isoform of Na+ channel (24). The
mechanisms, which cause the observed alterations in
INa voltage dependence, conductance, time to
peak activation and inactivation, remain unknown and must be resolved
in further studies.
Changes in ICa-L.
L-type Ca2+ current provides inward current that is
necessary to maintain the long plateau phase of the fish ventricular
cells, and it is also indispensable for the excitation-contraction
coupling. Density, voltage dependence of activation, kinetics of
activation, and decay of the ICa-L in freshly
isolated myocytes are similar as recorded previously in trout
ventricular and atrial myocytes (22, 25). After 2-3
days of culture, the density of ICa-L was
decreased, the voltage dependence was shifted to more positive potentials, and current decay was slower compared with freshly isolated
trout ventricular myocytes. The decrease in
ICa-L density was only transient as it recovered
to the initial control level after 8-10 days in culture. In
contrast, the kinetic changes persisted or increased after 8-10
days of culture.
Significant changes in ICa-L during culture have
been observed in a number of mammalian culture preparations.
ICa-L density shows great variability (increase,
decrease, or no change) between studies, which might indicate species
specificity, but could also be due to differences in culture methods or
recording conditions (see Ref. 16). A decrease in the
early phase of culture and the subsequent recovery of
ICa-L density to the initial control level of
freshly isolated myocytes has been observed in rat and rabbit
ventricular myocytes cultured in defined (6) or
nonsupplemented media (15). The behavior of
ICa-L in trout ventricular myocytes therefore
resembles, at least qualitatively, the ICa
density changes of mammalian cardiac myocytes in serum-free cultures,
suggesting that regulation of L-type Ca2+ channels may be
mediated by similar mechanisms in fish and mammalian hearts.
The changes in voltage dependence of steady-state activation and in the
rate of activation and decay are very similar for ICa-L and INa, which
suggest that they might have a common origin in the membrane
environment of these channels or in the channels themselves, e.g., due
to a change in surface charge. The voltage dependence of activation for
both ICa-L and INa was
shifted to more depolarized voltages, which could be due a net decrease
in extracellular or a net increase in intracellular negative surface charge. As sialic acids are intrinsic components in the carbohydrate moieties of the ion channel proteins and important anionic binding sites in the external surface of the SL, culture-dependent reduction in
the amount of sialic acids or other anionic binding sites could have
caused the changes in I-V relationship and current kinetics of ICa-L and INa.
Experiments where surface charges of the SL are screened with different
concentrations of external Ca2+ could possibly resolve this
issue. It should be noted, however, that changes in voltage dependence
of INa and ICa cannot
explain the large depression of INa and
ICa during culture, as both currents are much
smaller in cultured than freshly isolated cells both at strongly
positive voltages and at peak current.
Perspectives
In contrast to mammalian cardiac myocytes, the depression of
K+ and Na+ currents in trout ventricular
myocytes was not associated with any changes in capacitive membrane
area. Therefore, it seems unlikely that depression of these cation
currents is due to the specific loss of surface membrane in areas where
the channels are located. Transient downregulation of
ICa-L and appearance of another K+
conductance, probably IKr, during later phases
of culture are also inconsistent with the "membrane-loss"
hypothesis. Rather, the present results suggest that culture conditions
alter the expression of different cation channels without any major
changes in the morphology of the myocytes. SL of fish cardiac myocytes have variable amounts of membrane invaginations in the form of caveolae
that are thought to be precursors for t-tubules. The caveolae are
enriched in a variety of membrane-signaling molecules like G
protein-coupled receptors, inositol trisphosphate (IP3) receptors, Ca2+-ATPase, and protein kinase
C (for review, see Ref. 12). Although there is
currently no direct evidence for the presence of ion channels in
caveolae of cardiac myocytes in either fish or mammals, the possibility
remains that the reduction in the density of IK1 in this study was due to reduction in number of functional channels or
channel regulators in the caveolae of cultured trout ventricular myocytes. Localization and quantitation of K+ channels in
the SL are needed to test this possibility.
The present findings indicate that fish cardiac myocytes can be kept in
serum-free primary culture for at least 10 days at 17°C. The
electrophysiological characteristics of the myocytes change markedly
but in a well-predicted and consistent manner during culture. The
serum-free culture of the fish cardiac myocytes is a promising
preparation that can be used in the future to clarify how the
expression of sarcolemmal cation currents is regulated by temperature
and other environmental stresses.
| |
ACKNOWLEDGEMENTS |
Kontiolahti fish farm is gratefully acknowledged for supplying the trout.
| |
FOOTNOTES |
This study was supported by the Academy of Finland (project No. 63090).
Address for reprint requests and other correspondence: M. Vornanen, Univ. of Joensuu, Dept. of Biology, PO Box 111, 80101 Joensuu, Finland (E-mail:
matti.vornanen{at}joensuu.fi).
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
10.1152/ajpregu.00350.2001
Received 18 June 2001; accepted in final form 29 November 2001.
| |
REFERENCES |
1.
Barron, EA,
Markwald RR,
and
Nathan RD.
Localization of sialic acid at the surface of embryonic myocardial cells.
J Mol Cell Cardiol
14:
381-395,
1982[Web of Science][Medline].
2.
Bennett, E,
Urcan MS,
Tinkle SS,
Koszowski AG,
and
Levinson SR.
Contribution of sialic acid to the voltage dependence of sodium channel gating: a possible electrostatic mechanism.
J Gen Physiol
109:
327-343,
1997[Abstract/Free Full Text].
3.
Bérnardeau, A,
Hatem SN,
Rücker-Martin C,
Tessier S,
Dinaian S,
Samuel JL,
Coraboeuf E,
and
Mercardier JJ.
Primary culture of human atrial myocytes is associated with the appearance of structural and functional characteristics of immature myocardium.
J Mol Cell Cardiol
29:
1307-1320,
1997[Web of Science][Medline].
4.
Catterall, WA.
Structure and function of voltage-gated ion channels.
Ann Rev Physiol
64:
494-531,
1995.
5.
Christe, G.
Localization of K+ channels in the T-tubules of cardiomyocytes as suggested by the parallel decay of membrane capacitance, IK1 and IKATP during culture and by delayed IK1 response to barium.
J Mol Cell Cardiol
31:
2207-2213,
2000.
6.
Davidoff, AJ,
Prasad SK,
Berger HJ,
Springhorn JP,
Marsh JD,
Kelly RA,
and
Smith TW.
Adult rat ventricular myocytes cultured in a defined medium: phenotype and electromechanical function.
Am J Physiol Heart Circ Physiol
265:
H747-H754,
1993[Abstract/Free Full Text].
7.
Eickhorn, R,
Drägert C,
and
Antoni H.
Influence of cell isolation and recording technique on the voltage dependence of the fast cardiac sodium current of the rat.
J Mol Cell Cardiol
26:
1095-1108,
1994[Web of Science][Medline].
8.
Ellingsen, O,
Davidoff AJ,
Prasad SK,
Berger HJ,
Springhorn JP,
Marsh JD,
Kelly RA,
and
Smith TW.
Adult rat ventricular myocytes cultured in defined medium: phenotype and mechanical function.
Am J Physiol Heart Circ Physiol
265:
H747-H754,
1993.
9.
Feng, J,
Li GR,
Fermini B,
and
Nattel S.
Properties of sodium and potassium currents of cultured adult human atrial myocytes.
Am J Physiol Heart Circ Physiol
270:
H1676-H1686,
1996[Abstract/Free Full Text].
10.
Hamill, O,
Marty A,
Neher E,
Sakmann B,
and
Sigworth FJ.
Improved patch-clamp techniques for high-resolution current recording from cells and cell-free membrane patches.
Pflügers Arch
391:
85-100,
1981[Web of Science][Medline].
11.
Harwood, CL,
Howarth C,
Altringham JD,
and
White E.
Rate-dependent changes in cell shortening, intracellular Ca2+ levels and membrane potential in single, isolated rainbow trout (Oncorhynchus mykiss) ventricular myocytes.
J Exp Biol
203:
493-504,
2000[Abstract].
12.
Isshki, M,
and
Anderson RGW
Calcium signal transduction from caveolae.
Cell Calcium
26:
201-208,
1999[Web of Science][Medline].
13.
Jacobson, SL,
and
Piper HM.
Cell culture of adult cardiomyocytes as models of the myocardium.
J Mol Cell Cardiol
18:
661-678,
1986[Web of Science][Medline].
14.
Lipp, P,
Hüser J,
Pott L,
and
Niggli E.
Spatially non-uniform Ca2+ signals induced by the reduction of transverse tubules in citrate-loaded guinea-pig ventricular myocytes in culture.
J Physiol (Lond)
497:
589-597,
1996[Abstract/Free Full Text].
15.
Mitcheson, JS,
Hancox JC,
and
Levi AJ.
Action potentials, ion channel currents and transverse tubule density in adult rabbit ventricular myocytes maintained for 6 days in cell culture.
Pflügers Arch
431:
814-827,
1996[Web of Science][Medline].
16.
Mitcheson, JS,
Hancox JC,
and
Levi AJ.
Cultured adult cardiac myocytes: future applications, culture methods, morphological and electrophysiological properties.
Cardiovasc Res
39:
280-300,
1998[Abstract/Free Full Text].
17.
Pollack, PS,
Carson NL,
Nuss B,
and
Houser SR.
Mechanical properties of adult feline ventricular myocytes in culture.
Am J Physiol Heart Circ Physiol
260:
H234-H241,
1991[Abstract/Free Full Text].
18.
Santer, RM.
The organization of the sarcoplasmic reticulum in teleost ventricular myocardial cells.
Cell Tiss Res
151:
395-402,
1974[Web of Science][Medline].
19.
Schackow, TE,
Decker RS,
and
TenEick RE.
Electrophysiology of adult cat ventricular myocytes: changes during primary culture.
Am J Physiol Cell Physiol
268:
C1002-C1017,
1995[Abstract/Free Full Text].
20.
Schackow, TE,
Sheets MF,
Decker RS,
and
TenEick RE.
Alteration of the sodium current in cat cardiac ventricular myocytes during primary culture.
Am J Physiol Cell Physiol
268:
C993-C1001,
1995[Abstract/Free Full Text].
21.
Schmidt, JW,
and
Catterall WA.
Palmitylation, sulfation and glycosylation of the 
-subunit of the sodium channel: role of posttranslational modifications in channel assembly.
J Biol Chem
262:
13713-13723,
1987[Abstract/Free Full Text].
22.
Shiels, HA,
Vornanen M,
and
Farrell AP.
Temperature-dependence of L-type Ca2+ channel current in atrial myocytes from rainbow trout.
J Exp Biol
203:
2771-2880,
2000[Abstract].
23.
Veldkamp, MW,
deJonge B,
and
Van Ginneken ACG
Decreased inward rectifier current in adult rabbit ventricular myocytes maintained in primary culture: a single-channel study.
Cardiovasc Res
42:
424-433,
1999[Abstract/Free Full Text].
24.
Vornanen, M.
Sarcolemmal Ca influx through L-type Ca channels in ventricular myocytes of a teleost fish.
Am J Physiol Regulatory Integrative Comp Physiol
272:
R1432-R1440,
1997[Abstract/Free Full Text].
25.
Vornanen, M.
L-type Ca current in fish cardiac myocytes: effects of thermal acclimation and 
-adrenergic stimulation.
J Exp Biol
201:
533-547,
1998[Abstract/Free Full Text].
26.
Vornanen, M,
Ryökkynen A,
and
Nurmi A.
Temperature-dependent expression of sarcolemmal K+ currents in rainbow trout atrial and ventricular myocytes.
Am J Physiol Regulatory Integrative Comp Physiol
282:
R1191-R1199,
2002[Abstract/Free Full Text].
27.
Vornanen, M,
and
Tuomennoro J.
Effects of acute anoxia on heart function in crucian carp: importance of cholinergic and purinergic control.
Am J Physiol Regulatory Integrative Comp Physiol
277:
R465-R475,
1999[Abstract/Free Full Text].
28.
Wetzel, GT,
and
Klitzner TS.
Developmental cardiac electrophysiology. Recent advances in cellular physiology.
Cardiovasc Res
31:
E52-E60,
1996.
29.
Xie, LH,
Takano M,
and
Noma A.
Development of inwardly rectifying K+ channel family in rat ventricular myocytes.
Am J Physiol Heart Circ Physiol
272:
H1741-H1750,
1997[Abstract/Free Full Text].
Am J Physiol Regul Integr Comp Physiol 282(4):R1200-R1209
0363-6119/02 $5.00
Copyright © 2002 the American Physiological Society
TOP
ABSTRACT
INTRODUCTION
