Characteristics of rabbit ClC-2 current expressed in
Xenopus oocytes and its contribution
to volume regulation
Tetsushi
Furukawa1,
Takehiko
Ogura1,
Yoshifumi
Katayama1, and
Masayasu
Hiraoka2
Departments of 1 Autonomic
Physiology and 2 Cardiovascular
Diseases, Medical Research Institute, Tokyo Medical and Dental
University, Chiyoda-ku, Tokyo 101, Japan
 |
ABSTRACT |
In the
Xenopus oocyte heterologous expression
system, the electrophysiological characteristics of rabbit ClC-2
current and its contribution to volume regulation were examined.
Expressed currents on oocytes were recorded with a two-electrode
voltage-clamp technique. Oocyte volume was assessed by taking pictures
of oocytes with a magnification of ×40. Rabbit ClC-2 currents
exhibited inward rectification and had a halide anion permeability
sequence of Cl
Br
I
F
. ClC-2 currents were
inhibited by 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB),
diphenylamine-2-carboxylic acid (DPC), and anthracene-9-carboxylic acid
(9-AC), with a potency order of NPPB > DPC = 9-AC, but were resistant to stilbene disulfonates. These characteristics are similar
to those of rat ClC-2, suggesting rabbit ClC-2 as a counterpart of rat
ClC-2. During a 30-min perfusion with hyposmolar solution, current
amplitude at
160 mV and oocyte diameter were compared among
three groups: oocytes injected with distilled water, oocytes injected
with ClC-2 cRNA, and oocytes injected with ClC-2
NT cRNA (an open
channel mutant with NH2-terminal
truncation). Maximum inward current was largest in ClC-2
NT-injected
oocytes (
5.9 ± 0.4 µA), followed by ClC-2-injected oocytes
(
4.3 ± 0.6 µA), and smallest in water-injected oocytes
(
0.2 ± 0.2 µA), whereas the order of increase in oocyte
diameter was as follows: water-injected oocytes (9.0 ± 0.2%) > ClC-2-injected oocytes (5.3 ± 0.5%) > ClC-2
NT-injected oocytes (1.1 ± 0.2%). The findings that oocyte swelling was
smallest in oocytes with the largest expressed currents suggest that
ClC-2 currents expressed in Xenopus
oocytes appear to act for volume regulation when exposed to a
hyposmolar environment.
chloride channel; voltage clamp; cell swelling; ClC supergene
family
 |
INTRODUCTION |
THE UNDERSTANDING OF MOLECULAR aspects of
voltage-dependent Cl
channels has advanced remarkably since cloning of the
Cl
channel gene of
Torpedo marmorata (ClC-0) by the
hybrid-depletion approach (7). Since then, by homology screening or
polymerase chain reaction (PCR)-based screening, ten members of related
Cl
channels have been
cloned from various tissues in different species (8-10). This
Cl
channel supergene family
is now designated as the ClC family. Despite a growing body of
molecular data, understandings of endogenous counterparts and
physiological roles of the ClC family are limited to certain members of
the family. For instance, ClC-1 is responsible for a major
Cl
conductance in skeletal
muscle (19) and its abnormalities result in the muscle disorder
congenital myotonia (11, 18, 26). ClC-2 has unique features because
external hyposmolarity activates rat ClC-2 currents in
Xenopus oocytes, and the ClC-2 channel
has been suggested to act in some way in volume regulation under
hyposmolar conditions (5, 12, 20). The ClC-2 channel has
also been suggested to stabilize the relationship between the membrane
potential and the Cl
equilibrium potential in neurons (15-17).
A rabbit homologue of ClC-2 was cloned and was designated as ClC-2G
because this channel was first isolated from a rabbit gastric cDNA
library (13). Rat ClC-2 currents were examined using the two-electrode
voltage clamp of oocyte membrane (5, 12, 20), whereas rabbit ClC-2G
currents were characterized by voltage clamp of a lipid bilayer, in
which oocyte membrane injected with ClC-2G cRNA was incorporated (13).
Despite a high sequence homology, ion permeability differed between rat
ClC-2 currents and rabbit ClC-2G currents. Rat ClC-2 currents had a sequence of halide anion permeability of
Cl
Br
> I
, whereas rabbit ClC-2G
had a sequence of I
> Cl
. In addition, some
important characteristics found in rat ClC-2 currents were not fully
examined in rabbit ClC-2 currents or vice versa. Rat ClC-2 currents
were activated by external hyposmolarity, but effects of external
hyposmolarity on rabbit ClC-2 currents were not examined. Rabbit
ClC-2G currents were activated by application of the catalytic
subunit of protein kinase A (PKA), but the effects of PKA on rat ClC-2
were not completely clear. ClC-2G currents were activated by strong
external acidification (pH 3.0) and were suggested to be essential for
HCl secretion by the gastric parietal cells, but effects of external
acidification on rat ClC-2 currents were studied only in moderate
acidic conditions (pH 6.0) (12).
Previously, we isolated a clone that is identical to ClC-2G from the
rabbit heart cDNA library (ClC-2
) (3). During the screening of the
rabbit heart cDNA library, we also isolated a truncated form of
ClC-2
and we proposed the presence of an alternative splicing form
(ClC-2
). However, the new 5' sequence [35 base pairs
(bp)] is highly homologous to sequences at the very ends of many
sequenced cDNAs in different species, such as rice, and contains
repeats of palindromic sequence (12). Thus the truncated clone of
rabbit ClC-2 (ClC-2
) is likely to be an artifact of library
construction but not a product of alternative splicing (3). In that
study, expressed currents were not carefully separated from endogenous
Cl
currents of
Xenopus oocytes, and thus rabbit
ClC-2G (ClC-2
) currents were not fully characterized.
To assess whether rabbit ClC-2G (ClC-2
) is a tissue-different
variant due to alternative splicing of rat ClC-2 or just a counterpart
of rat ClC-2, it is imperative to fully characterize rabbit ClC-2
currents and compare them with those of rat ClC-2 currents. Thus, in
the present study, we further characterized rabbit ClC-2G (ClC-2
)
currents using two-electrode whole cell voltage clamp as used for
characterization of rat ClC-2 currents (5, 12, 20). Our data suggest
that rabbit ClC-2G (ClC-2
) currents had similar characteristics to
rat ClC-2 currents and ClC-2G (ClC-2
) appeared to be a counterpart
of rat ClC-2 in rabbit. Thus, in the present study, we use rabbit ClC-2
in place of rabbit ClC-2G (ClC-2
). Second, although it is well
documented that rat ClC-2 currents are activated by external
hyposmolarity (5, 12, 20), their possible role for cell physiology has
not been fully clarified. Thus we examined whether expression of ClC-2 in oocytes affected the volume regulation in response to external osmolarity changes.
 |
METHODS |
Molecular biology.
ClC-2 cDNA obtained from rabbit heart (3) was subcloned into pSPORT I
(GIBCO BRL, Rockville, MD). As reported for other members of the ClC
supergene family, including rat ClC-2,
Cl
currents could not be
expressed in Xenopus oocytes from the
original, putative full-length cDNA for rabbit (19, 20, 22). We had previously reported that outwardly rectifying
Cl
currents were expressed
by injection of cRNA transcribed from the original, putative
full-length cDNA for rabbit ClC-2 (3). However, this current is likely
to be an endogenous current present in oocytes but not an expressed
current of rabbit ClC-2. Thus DNA of the
Torpedo ClC-0 5' untranslated
region (83 bp) was synthesized and added to the first ATG of rabbit
ClC-2 by recombinant PCR methods (6). The sequence of the new chimeric
clone was verified by the dideoxynucleotide chain termination method
using a 373A DNA sequencing system (Perkin Elmer, Rockville, MD). In an
erratum to the previous study (3), we reported expression of inwardly rectifying Cl
currents by
injection of rabbit ClC-2 cRNA (3). Although not mentioned in this
erratum (3), in those experiments, DNA of the
Torpedo ClC-0 5' untranslated
region (83 bp) had been added to the first ATG of rabbit ClC-2. In
vitro methyl-capped complementary RNA (cRNA) was made using T7 RNA
polymerase (Stratagene, La Jolla, CA).
Oocyte handling and electrophysiology.
Xenopus oocyte preparation and
handling were carried out as described previously (24). In brief,
oocytes were removed from Xenopus
laevis (Hamamatsu Seibutsu, Hamamatsu, Japan) under
anesthesia, washed in Ca2+-free
OR-2 containing (in mM) 100 NaCl, 2 KCl, 1 MgCl2, 5 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES), and 5 tris(hydroxymethyl)aminomethane (Tris) (pH 7.6 with
HCl). Stage V and VI oocytes were defolliculated by treatment with 2 mg/ml collagenase (type IA; Worthington, Freehold, NJ) in
Ca2+-free OR-2 for ~30-60
min, washed extensively with
Ca2+-free OR-2 containing no
collagenase, and injected with ~30-50 ng of cRNA dissolved in 30 nl of diethylpyrocarbonate-treated distilled water. Injected oocytes
were incubated for ~4-6 days at ~12-18°C in modified
Barth's solution containing (in mM) 88 NaCl, 1 KCl, 2.4 NaHCO3, 15 Tris, 0.3 Ca(NO3)2,
0.4 CaCl2, and 0.8 MgSO4 and 100 µg/ml sodium
penicillin and 100 µg/ml streptomycin sulfate (pH 7.6 by HCl).
Membrane currents were recorded from oocytes with a two-electrode
voltage clamp using an amplifier (TEV-200; Dagan, Minneapolis, MN) at a
room temperature of ~24-26°C. Current-injecting and
potential-measuring electrodes had resistances of ~0.5-2.0 and
~1.0-3.0 M
, respectively, when filled with 3 M KCl. The bath
solution was connected to the ground via a low-resistance agarose
bridge containing 2% agarose in 3 M KCl. A second reference electrode
was used to avoid polarization errors. Junction potentials resulting
from solution changes were <2.5 mV in each experiment and were not
corrected. Current measurements were low-pass filtered at 0.5 kHz. Data
acquisition and analysis were done on an 80386-based microcomputer
using pCLAMP software and TL-1 analog-to-digital converter (Axon
Instruments, Foster City, CA). Oocytes were perfused continuously with
various external solutions. Each experiment began with perfusing a
modified ND96 solution with a mean osmolarity of 228 ± 11 mosM [solution 1 (isosmolar solution) in Table 1]. Oocytes were
kept in current-clamp mode for at least 5 min before switching to
voltage-clamp mode. Only oocytes exhibiting a resting membrane
potential negative to
30 mV were used. Oocytes were voltage
clamped at a holding potential of
30 or
10 mV in some
experiments, and 2-, 4-, or 5-s voltage steps were applied from
160 to +60 mV in 20-mV increments or from
120 to +60 mV
in 10-mV increments. Extracellular hyposmolarity was established by
perfusing a hyposmolar solution (109 ± 7 mosM), in which 96 mM
mannitol was omitted from the control isosmolar solution, leaving other
components the same (solution 2 in
Table 1), which was a slightly different method from others. In a study by Gründer et al. (5), 0.5× ND96 + 100 mM sucrose
was used as an isosmolar solution and hyposmolarity was achieved by
omission of 100 mM sucrose. We also used sucrose to balance external
osmolarity in a few experiments, but we obtained basically no different
findings between sucrose and mannitol used as an agent to balance
osmolarity. Osmolarity of the solution was measured using a vapor
pressure osmometer (model 5500; Wescor, Salt Lake City, UT). To
determine the ion permeability, we measured the reversal potential
(Erev) in
various external solutions. To determine the permeability for anions,
48 mM NaCl was replaced with equimolar sodium aspartate (solution 3), sodium glutamate
(solution 4), or sodium gluconate (solution 5). To determine the
permeability for cations, 48 mM NaCl was replaced with equimolar
choline chloride (solution 9) or KCl
(solution 10). To determine the
permeability for halide anions, 48 mM NaCl was replaced with equimolar
NaF (solution 8), NaBr
(solution 6), or NaI
(solution 7). To examine effects of extreme external acidification, 5 mM HEPES in external solution was
replaced with equimolar propionic acid for pH 4.6 (solution 11) or with citric acid
for pH 3.6 (solution 12), and pH was
adjusted with NaOH.
4,4'-Diisothiocyanostilbene-2,2'-disulfonic acid (DIDS;
Sigma, St. Louis, MO),
4-acetamido-4'-isothiocyanostilbene-2,2'-disulfonic acid
(SITS; Sigma), and 4,4'-dinitrostilbene-2,2'-disulfonic
acid (DNDS; Tokyo Kasei, Tokyo, Japan) were directly dissolved in
external test solutions just before experiments.
Anthracene-9-carboxylic acid (9-AC; Sigma) and
diphenylamine-2-carboxylic acid (DPC; Wako Pure Chemical, Osaka, Japan)
were prepared as 1 M stock solutions, 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB; Research Biochemical International, Natick, MA) as a 100 or 500 mM stock solution, and
forskolin (Wako Pure Chemical) as a 10 mM stock solution in dimethyl
sulfoxide (DMSO). They were stored at
20°C until needed and
were diluted in the external solution at final concentrations described
in the text. The final concentration of DMSO in each solution was
0.1%, which had no effect on the membrane currents of
Xenopus oocytes.
Photographs of oocytes were taken using a camera (model AFM; Nikon,
Ofuna, Japan) attached to the microscope with a magnification of
×40. Photographs of oocytes were copied with a magnification of
×2, and oocyte diameters were measured with a pair of dividers with a final magnification of ×80. To check the precision and accuracy of measurement of oocyte diameter, two researchers (Furukawa and Ogura) measured oocyte diameter independently and each researcher repeated measurements three times. Variations of measurement in each
researcher and variations of measurement between researchers never
exceeded 0.02 mm. Thus we consider the difference in oocyte diameter as
significant if the difference is >0.02 mm and had a
P value of <0.05.
All values are presented with n
(number of measurements in one oocyte) and
N (number of measurements in different
oocytes). Measurements were repeated three times
(n = 3) in the same oocyte if
possible. When the difference in measured values was <3 mV for
Erev, 5% of
measured values for current amplitude, and 0.02 mm for oocyte diameter,
measurement in that oocyte was considered to be stable and accurate and
included for further analysis. For the experiments in which external
osmolarity or pH was changed, current amplitude changed quickly from
time to time and repeatedly measured values varied substantially. Thus,
for these experiments, measurements were done singly
(n = 1) and stability of data was judged from continuous current recordings at a single voltage. Measured
values in different oocytes (N) were
averaged and are shown as means ± SE. One-way analysis of variance
followed by a Student's t-test was
used to test for significance (P < 0.05).
 |
RESULTS |
Current-voltage relationships and ion permeability.
In oocytes injected with rabbit ClC-2 cRNA, slowly activating inward
currents at negative potentials and small outward currents at positive
potentials were recorded (Fig.
1Aa).
For oocytes injected with distilled water, very small outward currents
but not inward currents were recorded (Fig.
1Ab), and, therefore, slowly
activating inward currents at negative potentials appeared to be
expressed currents due to injection of rabbit ClC-2 cRNA and outward
currents appeared to be endogenous currents of oocytes. The amplitude
of outward currents varied between different batches of oocytes, and we
arbitrarily decided to avoid oocytes in which current amplitude at +60
mV was >0.4 µA. Swelling-activated outward
Cl
currents and
hyperpolarization-activated inward
Cl
currents are
endogenously present in oocytes (1, 21). The former type of current was
decreased in amplitude after defolliculation and was almost
undetectable at 2 days after defolliculation (1). The latter type of
current shifted its voltage dependence to the negative potential after
defolliculation, and potential for activation threshold was more
negative than
160 mV at 4 days after defolliculation (21). Thus
we performed current measurements at least 4 days after
defolliculation. The current-voltage
(I-V)
curves constructed at the end of 5-s step pulses showed marked inward
rectification (Fig. 1B). Because
ClC-2 currents showed only a minor, if any, inactivation, to determine
ion selectivity through the rabbit ClC-2 channel, quasi-steady-state
I-V
curves were constructed at the end of 5-s step pulses in various ionic
solutions and the Erev values were
measured. Permeability for anions through rabbit ClC-2 channel was
studied by replacing 48 mM
Cl
with equimolar
aspartate, glutamate, or gluconate (Fig.
2A). The
Erev in
Cl
solution
(solution 1 in Table 1) was
15.1 ± 1.3 mV (n = 3, N = 10), which was shifted to +31.7 ± 4.2 mV (n = 3, N = 8) in aspartate solution
(solution 3), +33.0 ± 4.9 mV (n = 3, N = 9) in glutamate solution
(solution 4), and +35.8 ± 5.0 mV
(n = 3, N = 10) in gluconate solution
(solution 5).

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Fig. 1.
Current-voltage
(I-V)
relationships of rabbit ClC-2 currents.
A: representative membrane currents
recorded from an oocyte injected with cRNA of rabbit ClC-2
(a) and that injected with distilled
water (b). Membrane was held at
30 mV, step pulses for 5 s to various potentials between
160 and +60 mV in 20-mV intervals were applied, and recorded
currents were superimposed. B:
I-V
curves for membrane currents of oocytes injected with rabbit ClC-2 cRNA
and those injected with distilled water.
n, Number of measurements in 1 oocyte;
N, number of measurements in different
oocytes.
|
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Fig. 2.
Ion permeability through rabbit ClC-2 channels.
Aa: representative superimposed
currents of oocytes injected with rabbit ClC-2 in control external
Cl solution
(left), in aspartate solution
(middle), and in glutamate solution
(right). Membrane potential was held
at 30 mV, and step pulses for 4 s were applied to various
potentials between 120 and +60 mV in 10-mV intervals. To
minimize current flowing at holding potential and resulting changes in
intracellular ionic environment, holding potential was changed to
10 mV for glutamate and aspartate solutions.
Ab:
I-V
curves for rabbit ClC-2 currents in control external
Cl solution, in aspartate
solution, in glutamate solution, and in gluconate solution.
Ba: rabbit ClC-2 currents in control
external solution containing 48 mM
Na+ and 2 mM
K+
(left), in
Na+-free solution containing 0 mM
Na+ and 2 mM
K+
(middle), and in
high-K+ solution containing 0 mM
Na+ and 50 mM
K+
(right). Concentration of
Cl in these 3 test
solutions was identical (55.6 mM). Membrane potential was held at
30 mV, and step pulses for 4 s were applied to various
potentials between 120 and +60 mV in 10-mV intervals.
Bb:
I-V
curves of rabbit ClC-2 currents in control solution containing 48 mM
Na+ and 2 mM
K+ (48Na, 2K), in solution
containing 0 mM Na+ and 2 mM
K+ (0Na, 2K), and in solution
containing 0 mM Na+ and 50 mM
K+ (0Na, 50K).
|
|
To examine whether cations are also permeable through this channel,
extracellular Na+ concentration
and K+ concentration were changed,
and the Erev was
measured. When extracellular Na+
was decreased from 48 to 0 mM by replacing with equimolar choline or
when extracellular K+ was
increased from 2 to 50 mM by replacing choline with
K+, the
Erev was not
significantly changed (Fig. 2B). The
Erev was
13.2 ± 3.2 mV (n = 3, N = 9) in the control solution
(solution 1 in Table 1),
14.1 ± 5.0 mV (n = 3, N=9)
in the Na+-free solution
(solution 9), and
10.7 ± 2.9 mV (n = 3, N = 9) in the
high-K+ solution
(solution 10). Thus the rabbit ClC-2
channel has a permeability that is highly selective for anions over
cations.
We next examined permeability of halide anions through this channel by
replacing 48 mM extracellular
Cl
with equimolar
Br
,
I
, or
F
(Table 1 and Fig.
3). Even in the same solution, the
Erev varied between different batches of oocytes. To avoid influence of this variation on data analysis, the data were used when the
Erev could be
measured for all four solutions from the same oocyte. In this way, the
order of permeability sequence could be assessed accurately. The
Erev was
19.5 ± 2.2 mV (n = 3, N = 10) in
Cl
solution
(solution 1),
11.0 ± 2.6 mV (n = 3, N = 10) in
Br
solution
(solution 6), +12.0 ± 2.9 mV
(n = 3, N = 10) in
I
solution
(solution 7), and +15.8 ± 5.1 mV
(n = 3, N = 10) in F
solution
(solution 8). Thus the sequence of
halide anion permeability was
Cl
Br
I
F
. This selectivity
sequence is similar to that of rat ClC-2 currents and corresponds to
Eisenman sequence 4 (23).

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Fig. 3.
Permeability of halide anions through rabbit ClC-2 channels.
A: rabbit ClC-2 currents in external
Cl solution
(a), in
Br solution
(b), in
I solution
(c), and in
F solution
(d). Stimulation protocol used was
same as in Fig. 2A.
B:
I-V
curves in Cl solution, in
Br solution, in
I solution, and in
F solution.
|
|
Sensitivity to Cl
channel
blockers.
To understand sensitivity of ClC-2 currents to various
Cl
channel blockers, we
tested DIDS, SITS, DNDS, 9-AC, DPC, and NPPB. Rabbit ClC-2 currents
were insensitive to stilbene disulfonates. Amounts of 1 mM DIDS, 1 mM
SITS, or 10 mM DNDS did not inhibit inward currents [only data
for 10 mM DNDS are presented (Fig. 4)]. Application of 10 mM DNDS
slightly shifted the
I-V
curves in a depolarizing direction (Fig.
4Ba). In contrast,
9-AC, DPC, and NPPB inhibited rabbit ClC-2 currents (Fig. 4,
Ab and
Bb-d). Remaining current
amplitudes at
160 mV after drug application were expressed as a
fraction of the current level before drug application and are shown in
Fig. 4C. It was significantly smaller for 0.5 mM NPPB (0.49 ± 0.13) than for 1 mM DPC (0.72 ± 0.14; P < 0.01 vs. NPPB) or 1 mM 9-AC
(0.69 ± 0.07; P < 0.01 vs. NPPB) (P < 0.05). There was no significant
difference between 1 mM 9-AC and 1 mM DPC. Thus an order of inhibitory
potency was NPPB > DPC = 9-AC. The remaining current fraction for 10 mM DNDS assessed at
160 mV was >1 (1.23 ± 0.17). This was
mainly due to a positive shift of voltage dependence by DNDS. Figure
4D displays a dose-response curve for
NPPB. The data were fitted by a least-squares analysis using SigmaPlot
software (Jandel Scientific, Corte Madera, CA) according to the Hill
equation in the following form
|
(1)
|
where
I is current amplitude in the presence
of NPPB expressed as a fraction to the current amplitude in the absence
of NPPB, k is the NPPB concentration
([NPPB]) causing half-maximal inhibition, and
n is the Hill coefficient. The
obtained value for k was 0.98 ± 0.03 mM and that for n was 0.97 ± 0.03.

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Fig. 4.
Effects of various Cl
channel blockers. Aa: rabbit ClC-2
current recorded in the absence
(left) or presence
(right) of 10 mM
4,4'-dinitrostilbene-2,2'-disulfonic acid (DNDS).
Ab: rabbit ClC-2 currents recorded in
the absence (left) or presence
(right) of 1 mM
diphenylamine-2-carboxylic acid (DPC). Stimulation protocol used was
same as for Fig. 1. B:
I-V
curves of rabbit ClC-2 currents in the absence (filled symbols) or
presence (open symbols) of 10 mM DNDS
(a), 1 mM anthracene-9-carboxylic
acid (9-AC; b), 1 mM DPC
(c), or 0.1 mM
5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB;
d).
* P < 0.05 for current
amplitude between absence and presence of each drug.
C: residual current amplitude at
160 mV in the presence of drugs expressed as a fraction of
current amplitude in the absence of drugs.
* P < 0.05 between drugs.
D: dose-response relationships for
NPPB-induced inhibition of ClC-2 currents. Ordinate is current
amplitude expressed as a fraction of control current level, and
abscissa is a concentration of NPPB. A continuous line was obtained by
fitting Hill equation (Eq. 1) to the
data.
|
|
Effects of external acidification.
We examined the effects of perfusion with external solution of pH 4.6 and 3.6 on rabbit ClC-2 currents. In Fig.
5A,
membrane was held at
30 mV and 2-s hyperpolarizing pulses to
160 mV were applied every 15 s. When external solution was
switched to pH 4.6, current amplitude quickly increased. Thereafter,
current amplitude decreased slowly and reached the steady-state level at ~5 min. When external pH was returned to pH 7.6, current amplitude increased gradually and returned to the initial level at ~10 min of
washout. In a different series of experiments, step pulses to various
membrane potentials were applied in the control state at pH 7.6 (Fig.
5, Ba and
C), at 30 s (Fig. 5,
Bb and
C), or at 5 min (Fig. 5,
Bc and
C) after a start of
perfusion with solution of pH 4.6. Roughly 1 min was required to
complete one series of the step-pulse protocol, and, as one can see in
Fig. 5A, current amplitude changed
substantially within 1 min. Thus the effects of pH at the end of the
step-pulse protocol (e.g., +60 mV) might be different from those at its
beginning (e.g.,
160 mV). Despite this limitation, it can be
clearly seen that current amplitude was increased at 30 s after start
of perfusion with a solution of pH 4.6 and was decreased at 5 min. At
30 s after start of perfusion with solution of pH 4.6, the increase in
current amplitude was observed both in inward and outward currents.
Thus this initial increase in current amplitude did not result from a
shift of voltage dependency due to surface charge screening by protons.
In the external solution at pH <4.0, oocyte membrane became unstable and step pulses to extremely negative or positive potentials could not
be applied. Thus we applied 2-s step pulses to
120 and +40 mV
alternatively every 15 s and external pH was changed to 3.6 for 4 min
and returned to 7.6 (Fig. 5D). Both
inward currents at
120 mV and outward currents at +40 mV were
initially increased by external pH 3.6 and thereafter decreased. These
changes were observed in all four experiments performed.

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Fig. 5.
Effects of low pH in the external solution.
A: representative recording of rabbit
ClC-2 current when pH of external solution was changed from 7.6 to 4.6 and returned to 7.6. Membrane was held at 30 mV, and 2-s
hyperpolarizing pulses to 160 mV were applied every 15 s. In
this and Figs. 6, 7, and 9, inward currents are shown downward and
outward currents are shown upward. Top
trace: current recording at 30-mV holding
potential. Bottom trace: current
recording at 160-mV hyperpolarizing pulses. Times when recording
chamber was perfused with external solution of pH 7.6 or 4.6 are
indicated by bars on top of current recording.
B: superimposed ClC-2 currents in the
control solution of pH 7.6 (a), at
30 s after start of perfusion with solution of pH 4.6 (b), and at 5 min after start of
perfusion with solution of pH 4.6 (c). Stimulation protocol used was
the same as in Fig. 1. C:
I-V
relationships in pH 7.6 solution, at 30 s after start of perfusion with
solution of pH 4.6, and at 5 min after start of perfusion with solution
of pH 4.6. D: representative recording
of rabbit ClC-2 current when pH of external solution was changed from
7.6 to 3.6 and returned to 7.6. Membrane was held at 30 mV, and
2-s hyperpolarizing pulses to 120 or +40 mV were applied
alternately every 15 s. Top trace:
current recording at +40-mV depolarizing pulses.
Middle trace: current recording at
30-mV holding potential. Bottom
trace: current recording at 120-mV
hyperpolarizing pulses. Times when recording chamber was perfused with
external solution of pH 7.6 or pH 3.6 are indicated by bars on top of
current recording.
|
|
Effect of forskolin.
Channel activity of rabbit ClC-2 (ClC-2G) was enhanced by application
of the catalytic subunit of PKA to the cytosolic side of the lipid
bilayer (13), but effects of PKA on rat ClC-2 were not examined. In a
previous study, we reported that external application of forskolin
activated rabbit ClC-2 currents (3). However, in those studies,
currents enhanced by forskolin were outward Cl
currents that are now
believed to be endogenous currents. Thus we again examined effects of
forskolin to increase adenosine 3',5'-cyclic monophosphate
(cAMP) and activity of endogenous PKA on rabbit ClC-2 (Fig.
6). External application of 10 µM
forskolin did not affect ClC-2 current amplitude.

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Fig. 6.
Effects of forskolin. A:
representative recording of rabbit ClC-2 current when an oocyte was
perfused with solution containing 10 µM forskolin. Membrane was held
at 30 mV, and 2-s hyperpolarizing pulses to 120 mV were
applied every 5 s. Top trace: current
recording at 30-mV holding potential. Bottom
trace: current recording at 120-mV
hyperpolarizing pulses. At arrows marked with
a or
b, step pulses for 2 s to various
potentials between 160 and +60 mV in 20-mV intervals were
applied from a holding potential of 30 mV. Step-pulse protocols
were repeated 3 times to check the stability of recordings.
B: superimposed ClC-2 currents in the
control state (a) and in the
presence of 10 µM forskolin (b).
Stimulation protocol used was the same as for Fig. 1.
C:
I-V
relationships in the control solution and in the presence of 10 µM
forskolin.
|
|
Rabbit ClC-2 currents are sensitive to cell swelling.
Changing the superfusate from isosmolar solution (228 mosM) to
hyposmolar solution (109 mosM) resulted in an ~1-10% increase in oocyte diameter, which was dependent on both the types of cRNA injected and the stimulation protocol used. Similar to rat ClC-2 currents, the rabbit ClC-2 currents were small in isosmolar solution, but current amplitude was markedly increased by superfusion with hyposmolar solution (Fig. 7,
Ab-Cb). Reperfusion with
isosmolar solution completely or in some cases partially decreased
current amplitude. In oocytes injected with distilled water,
superfusion with hyposmolar solution did not significantly enhance
membrane currents (Fig. 7,
Aa-Ca).

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Fig. 7.
Effects of external hyposmolarity on rabbit ClC-2 currents.
A: representative continuous
recordings of rabbit ClC-2 currents for oocytes injected with distilled
water (a) and rabbit ClC-2 cRNA
(b) when external solution was
changed from isosmolar solution (228 mosM) to hyposmolar solution (109 mosM) and changed back to isosmolar solution. Membrane potential was
held at 30 mV, and 2-s hyperpolarizing pulses to 160 mV
were applied every 5 s. Top traces:
current recordings at a 30-mV holding potential.
Bottom traces: current recordings at
160-mV hyperpolarizing pulses. Times when oocytes were
externally superfused with hyposmolar solution are indicated by bars.
At points marked by arrows, 2-s step pulses were applied to various
potentials between 160 and +60 mV in 20-mV increments.
B: superimposed membrane currents
recorded from oocytes injected with distilled water
(a) and rabbit ClC-2 cRNA
(b) recorded in the control state
(left), in hyposmolar solution
(middle), and during washout
(right). Stimulation protocol used
was the same as in Fig. 1. C:
I-V
curves for water-injected oocytes
(a) and rabbit ClC-2 currents
(b) in the control state, in
hyposmolar solution, or during washout.
|
|
The enhancement of ClC-2 current amplitude in response to external
hyposmolarity was a slow phenomenon (see Fig.
7Ab), which seems to be compatible
with the hypothesis that slow cellular changes, probably oocyte
swelling, may be responsible for current enhancement (5). If this
hypothesis is correct, ClC-2 currents and oocyte swelling should change
with a similar time course in response to external osmotic changes, as
indeed they did. Representative photographs of oocytes at several time
points are displayed in Fig.
8Ab.
Oocyte diameter was increased in hyposmolar solution and was decreased
to the control level during washout of hyposmolar solution. Figure 8Bb
depicts the time course of changes in current amplitude at a
160-mV hyperpolarizing pulse and in oocyte diameter. In oocytes
injected with ClC-2 cRNA, increases in current amplitude became
apparent at ~10 min of superfusion with hyposmolar solution and
current amplitude increased more slowly after ~25 min (Fig. 8Bb). When external solution was
changed back to isosmolar solution, current amplitude
decreased relatively quickly and returned to the control level at ~20
min of washout (Fig. 8Bb). These
patterns were also observed from the increase in oocyte diameter during hyposmolar perfusion and decrease in oocyte diameter during washout (Fig. 8Bb). The correlation between
time course of changes in current amplitude and time course of changes
in oocyte diameter supports the hypothesis that the changes in oocyte
volume rather than the changes in extracellular osmolarity per se are
directly linked to current activation.

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Fig. 8.
Effects of external hyposmolarity on oocyte diameter.
A: representative pictures of oocytes
in the control state (left), at 30 min after superfusion with hyposmolar solution
(middle), and at 30 min after
washout (right) for oocytes injected
with distilled water (a), ClC-2 cRNA
(b), and ClC-2 NT cRNA
(c). Here and in Fig. 9, photographs
of oocytes are aligned so that 1 electrode is impaled from the left
side and the other from the right side. Diameter of oocytes was
measured from areas where electrodes were not impaled, namely, as a
distance between top and bottom edge of oocytes.
B: time courses of changes in current
amplitude and oocyte diameter for oocytes injected with distilled water
(a), with ClC-2 cRNA
(b), or with ClC-2 NT cRNA
(c). Membrane was held at 30
mV, and 2-s hyperpolarizing pulses to 160 mV were applied every
5 s. Current amplitudes at end of 160-mV hyperpolarizing pulse
and oocyte diameter were measured every 5 min and were plotted. Data
for oocyte diameter were expressed as percent increase compared with
the control value and plotted.
|
|
To assess whether expression of ClC-2 currents affects volume
regulation of oocytes, we monitored changes in current amplitude and
diameter in oocytes injected with distilled water (Fig. 8, Aa and
Bb), ClC-2 cRNA (Fig. 8,
Ab and
Bb), and ClC-2
NT cRNA (Fig. 8,
Ac and
Bc). ClC-2
NT is a clone obtained
during the course of screening, in which 76 amino acids were
artificially deleted from the NH2
terminus (3). Because the NH2
terminus of rat ClC-2 was demonstrated to be a critical region for
response to cell swelling and it expressed open channels (5), it was
worth examining volume changes in oocytes injected with ClC-2
NT cRNA in response to external hyposmolarity (Fig. 8,
Ac and
Bc). In the control state, there was
no significant difference in diameter between oocytes injected with
distilled water and those injected with ClC-2 cRNA: 1.13 ± 0.02 mm
(n = 3, N = 12) and 1.14 ± 0.01 mm
(n = 3, N = 27), respectively. The diameter of
oocytes injected with ClC-2
NT cRNA was significantly smaller (1.01 ± 0.02 mm; n = 3 and
N = 11) than in oocytes injected with
distilled water and with ClC-2 cRNA (P < 0.01), which may be explained by activation of inward current even
in the isosmolar solution in oocytes injected with ClC-2
NT cRNA.
Current amplitude in water-injected oocytes was not changed (
0.2 ± 0.2 µA in the control,
0.2 ± 0.2 µA at 30 min of
hyposmolarity, and
0.2 ± 0.2 µA at 30 min of washout), but
oocyte diameter was increased by external hyposmolarity and decreased
by washout (Fig. 8, Aa and
Ba). Current amplitude in oocytes
injected with ClC-2
NT cRNA was the largest in the isosmolar solution
(
5.8 ± 0.4 µA). Although current amplitude was not
affected by external hyposmolarity, it remained large (
5.9 ± 0.4 µA at 30 min of hyposmolarity). However, oocyte diameter was only
slightly increased by external hyposmolarity (Fig. 8,
Ac and
Bc). The maximum percent increase in
oocyte diameter attained in the hyposmolar solution was smallest for
oocytes injected with ClC-2
NT (1.1 ± 0.2%;
n = 1 and
N = 5), followed by oocytes injected
with ClC-2 (5.3 ± 0.5%; n = 1 and
N = 12), and was largest for oocytes
injected with distilled water (9.0 ± 0.2%;
n = 1 and
N = 7). Thus oocyte swelling by
hyposmolarity was smallest in oocytes with the largest inward
Cl
currents, suggesting
that expression of ClC-2 currents markedly influenced regulation of
oocyte volume.
We next tested whether the frequency of activation of ClC-2 currents
affected volume regulation of oocytes injected with ClC-2. In Fig.
9A,
representative time courses of changes in ClC-2 current amplitude are
displayed for when step pulses to
120 mV were applied every 15 s
and thereafter at every 5 s. External hyposmolarity increased current
amplitude at
120-mV hyperpolarizing pulses. The
hyposmolarity-induced increase in current amplitude was greater when
stimulated every 15 s than every 5 s. Figure
9B shows representative photographs of
oocytes whose membrane was hyperpolarized to
120 mV every 15 s
(Fig. 9, Ba and
Ca) and every 5 s (Fig.
Bb and
Cb). Oocyte diameter at 30 min of
hyposmolarity was greater with stimulation at 1/15 s than with
stimulation at 1/5 s. Increases in current amplitude (Fig.
9Ca) and oocyte diameter (Fig.
9Cb) by external hyposmolarity were
compared between two stimulation frequencies. Increases in current
amplitude (Fig. 9Ca) and oocyte
diameter (Fig. 9Cb) were
significantly greater by stimulation at 1/15 s than at 1/5 s.
Maximum current amplitude at
120-mV hyperpolarizing pulses
was
4.4 ± 0.2 µA (n = 3, N=5) at 1/15 s and
2.9 ± 0.3 µA
(n = 3, N = 5) at 1/5 s
(P < 0.01). The maximum increase in oocyte diameter was 7.8 ± 1.1% (n = 3, N = 5) at 1/15 s and 6.2 ± 0.3% at 1/5 s (n = 3, N = 5)
(P < 0.05).

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Fig. 9.
Effects of frequency of current activation on current enhancement and
oocyte swelling. A: representative
continuous recordings of ClC-2 currents induced by hyperpolarizing
pulses to 120 mV at every 15 s (left half of trace), followed by
at every 5 s (right half of trace). Times when oocytes were externally
superfused with hyposmolar solution are indicated by bars on top of
current recordings. B: representative
pictures of oocytes in the control state
(left), at 30 min after hyposmolar
perfusion (middle), and at 30 min
after washout (right) for
stimulation rates of 1/15 s (a) and
1/5 s (b).
C: time courses of changes in current
amplitude (a) and oocyte diameter
(b) by stimulation at 1/15 and 1/5
s. * P < 0.05 between
stimulation at 1/15 and 1/5 s.
|
|
 |
DISCUSSION |
Utilizing the Xenopus oocyte
heterologous expression system and two-electrode voltage-clamp
technique, we have characterized two important features of rabbit
ClC-2. First, expressed currents of rabbit ClC-2 had similar
electrophysiological features to rat ClC-2 currents, in terms of
I-V
relationships, ion selectivity, and sensitivity to
Cl
channel blockers. Rabbit
ClC-2 currents were activated by external hyposmolarity in a manner
similar to rat ClC-2 currents. Second, expression of rabbit ClC-2
regulated the swelling of oocytes in response to external
hyposmolarity. This regulatory action was dependent on activation of
ClC-2 currents.
The protein encoded by rabbit ClC-2 had, overall, 93% similarity and
82% identity to rat ClC-2 (13, 20). [We had reported that
identity of overall sequence of amino acid between rat ClC-2 and rabbit
ClC-2 was 81% in a previous study. This is because we used ClC-2
NT
(ClC-2
) for rabbit ClC-2. Identity between rat ClC-2 sequence and
full-length sequence of rabbit ClC-2 is 82% (3).] Despite the
high amino acid homology, reported ion permeability is different
between rat ClC-2 currents (5, 12, 20) and rabbit ClC-2 currents (13).
In addition, several important electrophysiological characteristics had
not been clarified between rat ClC-2 currents and rabbit ClC-2
currents. The sequence of halide anion permeability was
Cl
Br
> I
for rat ClC-2 currents
(20) and I
> Cl
for rabbit ClC-2G
currents (13). Rat ClC-2 currents were inwardly rectifying (20),
whereas rabbit ClC-2 currents had linear
I-V relationships of single-channel conductance and a higher open probability at a positive potential (+80 mV) compared with a negative potential (
80 mV) (13). However, voltage dependence of whole cell currents of rabbit ClC-2 had not been examined. Rat ClC-2 currents
were sensitive to carboxylate derivatives and were resistant to
stilbene disulfonates (11). Effects of
Cl
channel blockers on
rabbit ClC-2G currents have not been examined. These
electrophysiological characteristics are fundamental for channel
function and must be clarified to determine whether rabbit ClC-2G is a
tissue-different variant of rat ClC-2 due to alternative splicing or is
just its counterpart. In the present study, the I-V
relationships, ion permeability, and sensitivity to channel blockers of
rabbit ClC-2 currents were similar to those of rat ClC-2 currents.
Thus, when characteristics of expressed currents were examined in the
same Xenopus oocyte expression system,
rat ClC-2 and rabbit ClC-2 exhibited similar electrophysiological features.
Similarities between rat ClC-2 and rabbit ClC-2 are not restricted to
fundamental electrophysiological features. Rat ClC-2 currents were
sensitive to external osmolarity changes (5), but effects of external
osmolarity on rabbit ClC-2 currents had not been examined yet. We
showed that rabbit ClC-2 currents expressed in
Xenopus oocytes were activated by
external hyposmolarity, as were rat ClC-2 currents. Furthermore, rat
ClC-2 and rabbit ClC-2 were similar in terms of tissue distribution.
The rat ClC-2 transcript was present ubiquitously in almost all tissues
(20). Data from reverse transcription-PCR experiments in our previous
study (3) showed that rabbit ClC-2 transcript was present in every
tissue we tested. [In the previous study, we also reported the
presence of a transcript for ClC-2
NT (ClC-2
) in heart and brain.
However, we could not reproduce these data by repeated experiments
using different batches of
poly(A)+ RNA and a different set
of PCR primers, and thus this reverse transcription-PCR product may be
nonspecific products or products due to some experimental errors
(3).] Together, these data suggest that rabbit ClC-2 is a
counterpart of rat ClC-2 in rabbit and not a tissue-different variant.
Two intriguing modulations of rabbit ClC-2 were reported by Malinowska
et al. (13). First, rabbit ClC-2 currents were enhanced by application
of the catalytic subunit of PKA to the cytosolic side of membrane (13).
A consensus sequence for PKA phosphorylation is present in the
COOH-terminal cytoplasmic stretch of rabbit ClC-2, but not in rat
ClC-2, which was suggested to be functionally important (rat ClC-2 has
PKA sites in different positions) (13, 20). We also reported that
rabbit ClC-2 currents were enhanced by external application of
forskolin; however, the currents modulated by forskolin were outward
currents and are now believed to be endogenous currents of oocytes (3).
[This is also the case for different magnitudes of current
inhibition by 9-AC in the previous study (~60%; Ref. 3) and in the
present study (~31%).] In the present study, we found no
effects of forskolin on inwardly rectifying rabbit ClC-2 currents.
Expressed currents of the human homologue of ClC-2 that has a consensus
PKA site on the COOH-terminal cytoplasmic stretch were not affected by
manipulations to increase intracellular cAMP levels. One possible
explanation for this discrepancy is different experimental condition.
Malinowska et al. (13) applied the PKA catalytic subunit directly to
the cytoplasmic side of the membrane. Others (3, 12) applied forskolin
or increased intracellular cAMP, in which the local concentration of
PKA achieved in close vicinity to the channels might not be as high as
the level achieved by direct application of the PKA catalytic subunit.
Another possibility is that Malinowska et al. (13) and others studied
different classes of channels because a sequence of ion permeability
shows stark contrast
(Cl
< I
vs.
Cl
> I
). In human T84 cells, a
native Cl
current resembles
ClC-2, and it was reported that this current was inhibited by cAMP (2).
It may be possible that different expression systems may potentially
cause this discrepancy (the presence of PKA modulation of inward
Cl
in human T84 epithelial
cell and its absence of ClC-2 current expressed in
Xenopus oocytes). High local
concentrations of active PKA may be required for modulation of ClC-2 by
way of some anchoring protein, which is indeed the case for cardiac
L-type Ca2+ channels (4). The
Xenopus oocyte may not express the
anchoring protein of PKA and may not be an appropriate expression
system to test modulation by PKA because enhancement of L-type cardiac Ca2+ channel by PKA could be
reproduced in some mammalian cell lines (e.g., Chinese hamster ovary
cells) (25) but not in the Xenopus oocyte expression system (14). Thus the scenario that ClC-2 currents
would be modulated by PKA in native tissues, which could not be
reconstituted in Xenopus oocyte
heterologous expression system, is still possible. Further studies are
required to clarify this point.
Another important modulator of rabbit ClC-2 reported was external
acidification. When rabbit ClC-2 currents were studied by voltage clamp
of oocyte membrane incorporated into the lipid bilayer, current
amplitude was increased by extreme external acidification (pH 3.0)
(13). Thus it was suggested that rabbit ClC-2 (ClC-2G) was active at
low pH, which is essential for HCl secretion by the gastric parietal
cells. Recently, it was reported that rat ClC-2 currents were also
enhanced by moderate external acidification (pH 6.0), but effects of
extreme acidification on rat ClC-2 currents were not reported (12). In
the present study, we could lower pH of external solution to 3.6 with a
decent stability of current recordings. This difference may be due to
different experimental conditions. In the system of voltage clamp of
lipid bilayer, buffering of pH both in intra- and extracellular
solution is more rigid. However, in oocytes, low pH in extracellular
solutions may cause intracellular acidification, which may damage
oocytes and cause instability of current recording. Within the range
that we could study, external acidification initially enhanced and
thereafter inhibited rabbit ClC-2 currents. Because current enhancement
by external acidification was observed at the moment when low-pH solution reached the recording chamber, this action appeared to be a
direct action of external protons. On the other hand, inhibitory effects of low pH were attained gradually and reached steady-state levels after ~5 min of perfusion with low-pH solution and ~10 min
was required to wash out this inhibitory effect. Thus this inhibitory
effect is likely to be due to some secondary changes triggered by
external acidification, such as allosteric changes of channel proteins
or intracellular acidification. Further experiments are needed to
assess this intriguing finding. The latter possibility could especially
be tested in a system in which intracellular and extracellular pH could
be controlled more rigidly (e.g., inside-out patch experiment, lipid
bilayer voltage clamp).
The time course of changes in ClC-2 current amplitude correlated with
the time course of changes in oocyte diameter. The increase in current
amplitude was initially fast but became slower after 20 min of
superfusion with hyposmolar solution. A similar pattern was observed in
the time course of oocyte swelling. These findings support the
hypothesis, although indirectly, that changes in cell volume rather
than external osmolarity per se are linked to modulation of ClC-2
currents. Oocyte swelling evoked by external hyposmolarity was smallest
in oocytes in which expressed current was largest, whereas oocyte
swelling was largest in those oocytes in which current amplitude was
smallest. Thus ClC-2 currents appear to compensate oocyte swelling in
hyposmolar solution. Volume regulatory action appears to
be dependent on activity of ClC-2 currents. The more frequently
stimulation was applied to activate ClC-2 currents, the less oocyte
swelling was attained. These findings are consistent with the idea that
efflux of Cl
through
activated ClC-2 channels reduces intracellular osmolytes in association
with obligatory loss of water, resulting in reduction in oocyte volume.
We repeated measurement of oocyte diameter three times by two different
observers. Error range of measurement in the same observer and that
between two observers was 0.02 mm. A difference, then, >0.02 mm with
a P value <0.05 was considered to be
significant. Nevertheless, we have to admit that our method of
measurement of oocyte diameter still lacks precision and accuracy. Measurement of oocyte diameter with more accurate space resolution is
needed to confirm these intriguing conclusions.
In summary, our data indicate that rabbit ClC-2 appears to be a
counterpart of rat ClC-2, but not a tissue-different variant, because
rabbit ClC-2 currents have fundamentally similar electrophysiological characteristics to those of rat ClC-2 currents. Rabbit ClC-2 currents were enhanced by extracellular hyposmolarity in a similar way as rat
ClC-2 currents. For this finding, the rat ClC-2 channel was suggested
to be important for volume regulation in a condition such as tissue
edema. Our finding that the expression of ClC-2 diminished oocyte
swelling in response to external hyposmolarity that is dependent on
activation of ClC-2 currents appears to further confirm the implication
of ClC-2 for volume regulation.
 |
NOTE ADDED IN PROOF |
After acceptance of this paper, ClC-3 was reported to code a
volume-activated Cl
channel in the heart and probably
other tissues (D. Duan, C. Winter, S. Cowley, J. R. Hume, and B. Horowitz. Nature 390: 417-421, 1997).
 |
ACKNOWLEDGEMENTS |
We thank Dr. C. D. McCaig (University of Aberdeen) for critical reading
and for correcting the English and N. Sakaguchi for secretarial
assistance.
 |
FOOTNOTES |
This work was supported by a Grant-in-Aid for Scientific Research on
Priority Areas of Cardiac Development and Gene Regulation from the
Ministry of Education, Science, and Culture, Japan; by a grant from the
Japan Cardiovascular Research Foundation; by a Bayer Cardiovascular
Disease Research Scholarship; and by a grant from the Molecular
Cardiology Study Group, Japan.
Address for reprint requests: T. Furukawa, Dept. of Autonomic
Physiology, Medical Research Institute, Tokyo Medical and Dental Univ.,
2-3-10, Kandasurugadai, Chiyoda-ku, Tokyo 101, Japan.
Received 3 July 1997; accepted in final form 30 October 1997.
 |
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