1 Renal Unit, Massachusetts General Hospital East, Charlestown 02129; 2 Department of Medicine, Harvard Medical School, Boston 02115; and 3 Renal Section, Boston University Medical Center, Boston, Massachusetts 02118
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ABSTRACT |
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In this study,
patch-clamp techniques were applied to cultured neonatal mouse cardiac
myocytes (NMCM) to assess the contribution of cAMP stimulation to the
anion permeability in this cell model. Addition of either isoproterenol
or a cocktail to raise intracellular cAMP increased the whole cell
currents of NMCM. The cAMP-dependent conductance was largely anionic,
as determined under asymmetrical (low intracellular)
Cl conditions and symmetrical Cl
in the presence of various counterions, including Na+,
Mg2+, Cs+, and
N-methyl-D-glucamine. Furthermore, the
cAMP-stimulated conductance was also permeable to ATP. The
cAMP-activated currents were inhibited by diphenylamine-2-carboxylate,
glibenclamide, and an anti-cystic fibrosis transmembrane conductance
regulator (CFTR) monoclonal antibody. The anti-CFTR monoclonal antibody
failed, however, to inhibit an osmotically activated anion conductance,
indicating that CFTR is not linked to osmotically stimulated currents
in this cell model. Immunodetection studies of both neonatal mouse heart tissue and cultured NMCM revealed that CFTR is expressed in these
preparations. The implication of CFTR in the cAMP-stimulated Cl
- and ATP-permeable conductance was further
verified with NMCM of CFTR knockout mice
[cftr(
/
)] in which cAMP stimulation
was without effect on the whole cell currents. In addition, stimulation with protein kinase A and ATP induced Cl
-permeable
single-channel activity in excised, inside-out patches from control,
but not cftr(
/
) NMCM. The data in this report indicate that cAMP stimulation of NMCM activates an anion-permeable conductance with functional properties similar to those expected for
CFTR, thus suggesting that CFTR may be responsible for the cAMP-activated conductance. CFTR may thus contribute to the permeation and/or regulation of Cl
- and ATP-permeable pathways
in the developing heart.
cystic fibrosis; heart; adenosine 5'-triphosphate channels; chloride channels; adenosine 5'-triphosphate release
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INTRODUCTION |
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CYSTIC FIBROSIS IS A GENETIC DISEASE caused by
mutations of the gene that encodes for the cystic fibrosis
transmembrane conductance regulator (CFTR) (29, 30). CFTR is a member
of the superfamily of transport proteins known as the ATP-binding
cassette (ABC) transporters or traffic ATPase. ABC transporters have
the ability to transport various substrates (13), and CFTR, in
particular, is associated with the cAMP-dependent movement of both
Cl and ATP (4, 28).
Recent studies indicate that the cAMP-activated Cl
conductance of cardiac cells may be associated with the expression of
CFTR (16, 17, 22). CFTR is present in various mammalian heart preparations including rabbit (39) and guinea pig (22). However, studies have suggested that adult rat and mouse cardiac myocytes lack a
cAMP-activated Cl
conductance, consistent with the
absence of a functional CFTR (8, 18). Similar findings have been
reported in adult human cardiac myocyte preparations (24). Recently, we
and others have provided evidence suggesting the presence of a
functional CFTR in the neonatal rat cardiac myocyte (40, 42, 43). This
raises the possibility of a developmentally controlled aspect of CFTR expression in different preparations.
In this report, primary cultures of neonatal mouse cardiac myocytes
(NMCM) from control mice, cftr gene knockout
[cftr(/
)] mice, and mice heterozygous
for the disrupted cftr gene
[cftr(+/
)] were used to assess the effect of
cAMP stimulation in the activation of anion-selective whole cell
conductances. By applying voltage-clamp techniques, we functionally
characterized a cAMP-inducible, time-independent, and
Cl
- and ATP-permeable conductance, which was absent
in NMCM obtained from cftr(
/
) mice. The
cAMP-activated anion currents were inhibited by
diphenylamine-2-carboxylate (DPC), glibenclamide, and anti-CFTR antibodies. The results in this report are consistent with the presence
of a functional cardiac CFTR in NMCM, which may play a relevant role in
the developing heart.
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MATERIALS AND METHODS |
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Primary cultures of NMCM. Primary cultures of NMCM were obtained with procedural modifications to a commercial isolation kit originally developed for neonatal rat cardiac myocytes (Worthington Biochemicals, Freehold, NJ). Pregnant control mice (C57BL/6 +/+, P100) were a kind gift of Drs. Richard L. Sidman and Aizhong Li (New England Regional Primate Research Center, Dept. of Neurology, Harvard Medical School). Briefly, beating hearts were harvested from <24-h-old neonatal mice and immediately placed in a Ca2+- and Mg2+-free Hanks' balanced salt solution (HBSS; Worthington). The hearts were minced and subjected to trypsin digestion (100 µg/ml in HBSS) for 16-18 h at 4°C. Trypsin digestion was stopped by addition of trypsin inhibitor (Worthington). Further collagenase digestion (type II collagenase, 150 U/ml; Worthington) was conducted at 37°C on a shaking bath for 45 min. Cell clumps were filtered through a 70-µm nylon filter, centrifuged, and washed with fresh Leibovitz L-15 medium. Cell pellets were resuspended in Ham's F-10 medium with L-glutamine (BioWhittaker, Walkersville, MD) also containing 5% bovine serum and 10% horse serum (BioWhittaker). Cells were seeded onto glass coverslips and allowed to grow at 37°C in an incubator gassed with 5% CO2. Cells attached and spread after 1 day in culture and were usually used for the patch-clamping experiments within 1 wk with no electrical differences in the ionic conductances studied.
CFTR knockout mice. Adult mice heterozygous for the disrupted cftr gene were obtained from Jackson Laboratories (Cftrtm1Unc; Bar Harbor, ME). These mice were originally generated by targeted gene disruption, involving insertion of the neomycin gene into the murine cftr gene (36). Mice homozygous for the disrupted gene display many features common to human cystic fibrosis patients, including failure to thrive, alteration of mucous and serous glands, and obstruction of glandlike structures (36). The mice were mated, and offspring were genotyped for the presence of the functional and disrupted cftr genes, and their hearts harvested and processed, <24 h after delivery, as described above.
PCR screening for the cftr gene. The presence of either wild-type or the disrupted cftr genes was determined by PCR of genomic DNA isolated from the mouse tails as previously described (15). PCR was conducted with primers (Ransom Hill Bioscience, Ramona, CA) specific for either the wild-type or mutant cftr genes. The primers were 5'-TGA ACC TTA GTC CTA TGT TGC C-3' (common), 5'-TCG AAT TCG CCA ATG ACA AGA C-3' (mutant), and 5'-CTT TGA TAG TAC CCG GCA TAA TC-3' (wild type), which produce either a 500 or 300 bp product for the mutant and wild-type alleles, respectively. Amplification of 100 ng DNA was performed over 30 cycles in a thermal cycler (480; Perkin-Elmer, Norwalk, CT). The primer sequences and reaction conditions were those specified by Jackson Laboratories, except that 3 mM MgCl2 was used in the reaction buffer to amplify the mutant sequence instead of the recommended 1.5 mM. The PCR amplification products were separated by electrophoresis in a 1.2% agarose gel.
Immunoprecipitation of CFTR. To assess the presence of
wild-type CFTR in cultured myocytes, pooled batches of NMCM were grown as indicated in 25-cm2 (50-ml) culture flasks. WT-1 cells
transfected to overexpress CFTR and used as a positive control (4) were
grown on 10-cm petri dishes. Cells (NMCM and WT-1) were washed (2 times) with PBS then scraped with a rubber policeman. Cell pellets were
obtained by differential centrifugation. Harvested cells in culture
were resuspended in immunoprecipitation (IP) buffer [1% Triton
X-100, 0.5% Nonidet P-40, 150 mM NaCl, 10 mM
Tris · HCl pH 7.5, 1 mM EGTA, 1 mM EDTA, 0.25 mM
NaVO4, 10 µg/ml phenylmethylsulfonyl fluoride (PMSF), and
10 µg/ml aprotinin] and then sonicated on ice. Sonicated
samples were centrifuged at 10,000 g for 10 min at 4°C. The
supernatant was transferred and then incubated for 16-18 h with a
monoclonal antibody (MAb; 2 µg antibody · mg
protein1 · ml IP
buffer
1) either directed against the COOH terminus
(MAb 24-1; Genzyme) or directed against the R-domain of CFTR (MAb
13-1; Genzyme). Immobilized protein A agarose was added to the
solution during the final 2 h of incubation. An agarose pellet was
obtained by centrifugation in a table microcentrifuge and washed three
times with ice-cold IP buffer (1 ml). Agarose samples containing 300 µg total protein were mixed with sample buffer and heated to 60°C for 5 min before loading. After being separated by 6% SDS-PAGE, samples were transferred to a polyvinylidene fluoride
membrane. The membrane was blocked with 5% nonfat dry milk for 1 h at
room temperature and then probed with the anti-CFTR antibody (MAb
24-1; Genzyme) for 18 h at 4°C, followed by a horseradish
peroxidase-based enzyme-linked chemiluminescence system.
The presence of CFTR was also determined directly from myocardial tissue obtained from neonatal mice. Hearts were harvested from neonatal mice <24 h old and immediately placed in Ca2+- and Mg2+-free PBS containing 20 µM PMSF. The hearts were minced, homogenized, then processed as described above. Tissue samples contained 1 mg total protein.
Whole cell currents. Currents and command voltages were
obtained and driven as previously described (28). Actual currents and
step potentials were obtained and driven with a Dagan 3900 (Dagan,
Minneapolis, MN). Signals were obtained at 4 kHz, filtered at 1.5 kHz
with an eight-pole Bessel filter (Frequency Devices, Haverhill, MA),
and stored in a hard disk of a personal computer to be analyzed with
pCLAMP 6.0.3 (Axon Instruments, Foster City, CA). The patch pipette
contained (in mM) either 30 KCl, 120 potassium aspartate, 1.0 KH2PO4, 1.0 MgCl2, 10 NaCl, 5.0 ATP, 5.0 EGTA, and 5.0 HEPES, pH adjusted to 7.4 with KOH, or 140 NaCl,
5.0 KCl, 5.0 ATP, 1.0 MgCl2, and 10 HEPES, pH adjusted to
7.4 with NaOH. The bathing solution contained (in mM) 140 NaCl, 5.0 KCl, 1.0 CaCl2, 1.0 MgCl2, and 10 HEPES, pH
adjusted to 7.4 with NaOH. In some experiments, whole cell currents
were measured in symmetrical Cl with
Cs+, N-methyl-D-glucamine (NMG), or
Mg2+ as the counterion where both the bath and pipette
solutions contained (in mM) either 140 CsCl, 140 NMG-Cl, or 70 MgCl2, along with 10 HEPES (pH adjusted to 7.4 with NMG).
CaCl2 (1 mM) was added to the bath solution to facilitate
the formation of the seals between the pipette and cell membrane.
Asymmetrical ATP/Cl
currents were assessed by
filling the patch pipette with a solution containing MgATP (100 mM,
adjusted to pH 7.4 with NMG) and then backfilled with one of the
solutions above, as previously described (1, 28). Whole cell
current-voltage relationships were obtained by applying 20-mV voltage
steps for 500 ms between +100 and
100 mV from a holding
potential of 0 mV. The whole cell conductance was calculated from
currents measured at 490 ms after applying the voltage steps. This
voltage protocol effectively eliminates the contribution of
time-dependent Na+ and Ca2+ currents.
NMCM beat spontaneously, in agreement with other neonatal mouse cardiac
myocyte preparations (23). Only single, spontaneously beating cells
were chosen for electrophysiological recordings. However, in the few
cases (~10%) where a cell in a cluster was selected instead, the
whole cell currents obtained were consistent with those of single cells
under the same experimental conditions, thus ruling out possible
electrical coupling with neighboring cells. After the whole cell patch
was obtained, the maximum repolarization potential (maximum diastolic
potential) for the cardiac myocytes still beating was 61.3 ± 6.1 mV, whereas the overshoot of the action potential was 35.5 ± 5.1 mV (n = 5). The average beating rate was 54 ± 11 beats per
min (n = 5). The resting potential measured using the
zero-current-clamp technique was
76.3 ± 1.0 mV
(n = 6). NMCM are largely round in shape, and the whole cell capacitance of wild-type and cftr(
/
) were similar
[42.5 ± 6.9 pF, n = 11, vs. 31.8 ± 10.0 pF, n = 6, for wild-type and cftr(
/
) NMCM,
respectively, P < 0.3]; therefore, no correction for
whole cell current was conducted.
Single-channel currents. The excised, inside-out patch-clamp
configuration was carried out as previously described (10). Currents
and command voltages were obtained and driven with a Dagan 3900 with
the use of sampling and filtering frequencies as described for the
whole cell experiments. Data were stored in a hard disk of a personal
computer and analyzed with pCLAMP 6.0.3. Data were further filtered at
200 Hz for display purposes. These experiments were performed in the
symmetrical Cl solutions described previously. Data
from the excised, inside-out patches with upward and downward
deflections indicating the channel open states for positive and
negative holding potentials, respectively, were obtained between +100
and
100 mV.
ATP release assay. ATP release from NMCM was assessed with the luciferin-luciferase assay using a monolight 2010 luminometer (Analytical Luminescence Laboratory, San Diego, CA) and methods previously described (1, 26). Briefly, extracellular and total cellular ATP were measured from confluent NMCM grown on glass coverslips. At the time of the experiment, the coverslips were placed in 12 × 75 mm plastic cuvettes containing 0.1 ml of the luciferin-luciferase assay mix and 0.5 ml of a Ca2+-free solution containing (in mM) 140 NaCl, 5 KCl, 0.8 MgCl2, and 10 HEPES, pH 7.4. The purified luciferin-luciferase solution (2005/2003; Analytical Luminescence Laboratory) contained 3 µg/ml luciferase and 400 µM luciferin in 25 mM HEPES buffer (pH 7.75; 2550; Analytical Luminescence Laboratory). The HEPES buffer also contained 10 mM MgCl2, a cofactor of the luciferin-luciferase reaction. The final pH of the mixture was 7.40. Photon release was continuously measured in a luminometer (MonoLight 2010; Analytical Luminescence Laboratory). Coverslips were placed in a plastic cuvette after washing the "contaminating" culture medium and were held vertically with microclips (Roboz Surgical Instruments, Rockville, MD) (26). The ATP release was followed by the photon release of the luciferin-luciferase assay for ~2 min. To determine the amount of ATP released from cells, known concentrations of ATP in solution were also measured to construct a calibration curve.
To activate the cAMP-dependent stimulatory pathway of NMCM, cells were incubated with cholera toxin (CT; 1 µg/ml, Sigma) for 12 h, which does not interfere with the luciferin-luciferase assay (26). Neither cell volume nor cell count changed significantly after CT treatment. Both control and CT-treated cells excluded trypan blue, an indication that cell permeability was not impaired by the experimental conditions imposed. This was further supported by the ability of CT-treated cells to spontaneously beat.
Drugs and chemicals. The cAMP-stimulatory cocktail contained 8-bromoadenosine 3',5'-cyclic monophosphate (8-Br-cAMP; 500 µM; Sigma Chemical, St. Louis, MO), IBMX (200 µM; Sigma), and forskolin (10 µM, Sigma). The cAMP analog 8-Br-cAMP was used from a 25 mM stock in a 1:1 (vol/vol) ethanol/DMSO solution. Forskolin was used from a 100% ethanol, 50 mM stock solution. The phosphodiesterase inhibitor IBMX was used from a 20 mM solution in ethanol. Use of this cocktail, which results in a maximum concentration of 1.6% ethanol and 0.3% DMSO in the bathing solution, was without effect on any NMCM electrical parameters (data not shown). The CFTR channel blocker DPC (Fluka Chemical, Ronkonkoma, NY) was kept in a 100-fold stock solution (20 mM) in 50% water and ethanol. Glibenclamide (Research Biochemicals International, Natick, MA) was kept in a 10 mM stock solution in 100% DMSO. Isoproterenol (Sigma) was freshly prepared in water (0.1 mM) and used at a final concentration of 1 µM. The monoclonal antibody raised against the R-domain of CFTR (MAb 13-1; Genzyme) was directly diluted 1:100 times in the intracellular solution from a stock solution (292 µg/ml).
Statistical analysis. Average data values were expressed as means ± SE for each group tested. Statistical significance was obtained by Student's t-test for paired data (35) unless otherwise specified in the results. Data were considered significantly different when P < 0.05.
The perm-selectivity ratio
PATP/PCl was calculated with a
derivation of the Goldman-Hodgkin-Katz equation (14) from the cAMP-stimulated Cl and ATP currents obtained under
asymmetrical conditions, such that
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RESULTS |
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Whole cell currents of NMCM in low intracellular
Cl.
To assess the presence of a cAMP-dependent anion conductance in
neonatal cardiac myocytes, whole cell Cl
currents
were recorded with the voltage-clamp technique (10) under physiological
conditions, namely, the presence of low intracellular (42 mM) and high
extracellular (149 mM) Cl
concentrations. Addition
of isoproterenol (1 µM) induced a 499% increase in the whole cell
conductance for positive holding potentials (1.36 ± 0.23 vs. 8.14 ± 0.45 nS/cell, n = 4, P < 0.001; Fig.
1A) and a 161%
increase in whole cell conductance for negative holding potentials
(1.09 ± 0.16 vs. 2.84 ± 0.63 nS/cell, n = 4, P < 0.025; Fig. 1A). Although the isoproterenol induced whole cell
currents in asymmetrical Cl
rectified
(785 ± 66 pA/cell, n = 3, vs.
251 ± 46 pA/cell,
n = 4, P < 0.005, at ±100 mV, respectively; Fig.
1A) in the direction expected for activation of an
anion-permeable conductance, the reversal potential
(Er) following isoproterenol was only
23.0 ± 1.0 mV (n = 4), thus different from the predicted
Er of
33 mV if the currents were carried
exclusively by Cl
, but consistent with previous
reports (9, 11).
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Whole cell currents of NMCM in symmetrical Cl.
To further assess the anion-dependence of the cAMP-activated
conductances, whole cell currents were assessed in the presence of
different Cl
counterions. Addition of the
cAMP-stimulatory cocktail induced an increase in highly linear whole
cell currents with various salts. In symmetrical NaCl, addition of the
cAMP-stimulatory cocktail induced a 2,421% increase (0.33 ± 0.11 vs.
8.32 ± 2.31 nS/cell, n = 8, P < 0.005; Fig.
2A) in the linear whole cell
conductance. Similar findings were obtained in symmetrical NMG-Cl (0.28 ± 0.17 vs. 7.97 ± 2.38 nS/cell, n = 4, P < 0.02;
Fig. 2B), MgCl2 (3.45 ± 1.48 nS/cell, n = 8, vs. 15.0 ± 3.60 nS/cell, n = 6, P < 0.02; Fig.
2C), and CsCl (1.33 ± 0.24 vs. 8.74 ± 1.60 nS/cell,
n = 6, P < 0.005; Fig. 2D). The
cAMP-stimulated whole cell currents in symmetrical NaCl were inhibited
with DPC (400 µM, n = 3, P < 0.05, Fig
3), but not DIDS (400 µM, P < 0.8, n = 3, Fig. 3).
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Whole cell currents of NMCM in asymmetrical
ATP/Cl.
Expression of CFTR is associated with a cAMP-activated pathway that is
permeable to both Cl
and ATP (4, 28). Thus, to
assess the presence of ATP-permeable pathways in mouse cardiac
myocytes, whole cell currents were obtained in the presence of
intracellular MgATP (100 mM) and bathing Cl
(149 mM). Addition of the cAMP-stimulatory cocktail induced a 1,058% (0.43 ± 0.05 vs. 5.41 ± 0.40 nS/cell, n = 5, P < 0.001; Fig. 8) stimulation
of highly linear whole cell currents, indicative of an
electrodiffusional pathway permeable to both Cl
and
ATP. A 38-mV shift (54 ± 8 vs. 16 ± 5 mV, n = 5, P < 0.001) in the Er was observed after
cAMP stimulation, in agreement with activation of an anionic conductive
pathway. The change in Er was also consistent with
a perm-selectivity ratio for ATP/Cl
of 0.42 or 0.37 for zATP =
2 (MgATP) or
4 (free ATP),
respectively. Thus the ATP/Cl
perm-selectivity ratio
of the cAMP-activated anion-selective pathway of NMCM was in agreement
with our previous report on cells expressing the epithelial isoform of
CFTR (28) and with our previous report on CFTR reconstituted into lipid
bilayers (4). Both cAMP-stimulated ATP and Cl
currents were simultaneously inhibited by DPC (65.7%; Fig. 8), thus
further suggesting a permeation of Cl
and ATP
through the same anionic pathway. Inhibition was slightly higher at
positive potentials, however, suggesting a competition between
Cl
and ATP (Fig. 8B) for the blocking effect
of DPC. Interestingly, a similar effect was also observed in the
presence of low Cl
(Fig. 1B), where the
change in slope conductance was 76% and 57% for positive and negative
potentials, respectively.
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Whole cell currents in NMCM from CFTR knockout mice in symmetrical
Cl.
To further assess the functional role of CFTR in the cAMP-activated
Cl
current of NMCM, whole cell currents were also
obtained in NMCM from mice lacking the gene for CFTR (36). Whole cell
currents from cftr(
/
) NMCM were obtained in
symmetrical Cl
under conditions similar to those
reported above. In the presence of the NaCl solution on both sides of
the membrane, the basal whole cell conductance was 1.18 ± 0.20 nS/cell (n = 6). Addition of the cAMP-stimulatory cocktail was
without effect on the whole cell conductance (1.63 ± 0.32 nS/cell,
n = 6, P < 0.4; Fig.
9A).
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Whole cell currents in NMCM from CFTR knockout mice in asymmetrical
ATP/Cl.
To further assess the potential role of cardiac CFTR in the regulation
of an ATP-permeable pathway, cAMP-activated whole cell currents were
also obtained under asymmetrical ATP/Cl
conditions
in both cftr(+/
) and cftr(
/
) NMCM.
The cAMP-stimulated whole cell currents from heterozygous cardiac
myocytes were significantly larger than those from CFTR-knockout
myocytes (7.94 ± 0.1 nS/cell, n = 2, vs. 0.34 ± 0.16 nS/cell, n = 4, P < 0.001 by unpaired t-test, data
not shown). The data are consistent with a role of CFTR expression in
the cAMP-activated Cl
and ATP currents.
Single-channel currents from control NMCM in symmetrical
Cl.
The nature of the electrodiffusional pathway for Cl
was further investigated at the single-channel level. Excised patches from control NMCM were obtained in either symmetrical NaCl (140 mM,
n = 5), or MgCl2 (70 mM, n = 3)
solutions. Channel activity was observed after addition of MgATP (5 mM)
and protein kinase A (PKA; 50 ng/ml) in four of five experiments in
NaCl and three of three experiments in MgCl2, but no
channel activity was observed with either MgATP or PKA alone. The
single-channel conductance of the PKA-activated channels was 17.7 ± 2.2 pS/cell (n = 7).
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Single-channel currents from cftr(/
) NMCM in
symmetrical Cl
.
The presence of Cl
channel currents in
cftr(
/
) NMCM was also sought. In symmetrical
MgCl2 (70 mM), addition of MgATP (5 mM) and PKA (50 ng/ml)
to the cytosolic side of the patch failed to stimulate channel activity
in five of five patches. However, MgATP and PKA stimulated
single-channel activity in six of seven patches obtained from the
cftr(+/
) littermate NMCM (Fig.
12). These channels had a conductance of
11.4 ± 2.0 pS/cell (n = 6) that was statistically similar to
the single-channel conductance observed in the control NMCM (P < 0.1).
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DISCUSSION |
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CFTR has been recently observed in the mammalian heart (17, 22). The
pattern of expression of cardiac CFTR, as well as its physiological
significance, however, are the subject of current attention.
Interesting tissue and region-specific differences have been reported.
Rabbit (39) and guinea pig (3, 11, 12) cardiac tissues express sizable
cAMP-activated Cl currents that are associated with
a functional CFTR. Conversely, the lack of cAMP-dependent
Cl
currents has been attributed to the absence of
CFTR in both dog (38) and human hearts (24). The expression of cardiac
CFTR may also be region specific, because ventricular but not atrial rabbit cells seem to express this channel protein (16, 39). Adult rat
cardiac myocytes also lack a cAMP-dependent Cl
conductance consistent with the absence of a functional CFTR (8).
However, more recent evidence suggests that neonatal rat cardiac
myocytes express a cAMP-dependent Cl
conductance
identified as CFTR by Western blotting and functional assays (40). This
may represent a developmentally regulated pattern of expression.
In the present study, we addressed the issue as to whether CFTR
expression is associated with a cAMP-dependent anion conductance in the
neonatal mouse heart. Although osmotic- and purinergic-regulated (18)
Cl channels have been reported in this model, the
presence of CFTR has been, heretofore, largely unknown. The presence of
a cAMP-dependent anion conductance in NMCM was assessed by addition of
either isoproterenol or a cAMP-stimulatory cocktail. Both
cAMP-stimulating maneuvers increased rectifying currents under
asymmetrical Cl
conditions (low intracellular
Cl
) consistent with a Cl
conductance, which was also DPC inhibitable. The shift in the Er after cAMP stimulation, however, was not
unequivocally consistent with the sole activation of an anionic
conductance. Nevertheless, the observed Er was in
agreement with the report by Harvey et al. (11), where the
Er was
23.4 mV, with 42 mM
Cl
inside and 151 mM Cl
outside
the cell (the predicted Er in their study was
33 mV). Ehara and Ishihara (9) also reported on an
epinephrine-dependent Cl
conductance in guinea pig
ventricular myocytes, where discrepancies between the predicted and
measured Er were observed. The conditions in both
of these previous studies favored the possibility of contaminating cation conductances (9, 11) that could partially account for the change
in Er in the present study. Nevertheless, in
symmetrical Cl
with either Na+, NMG,
Cs+, or Mg2+ salts, cAMP stimulation induced a
highly linear and time-independent whole cell current, thus further
indicating that the cAMP-stimulated whole cell current was largely
accounted for by Cl
movement.
The cAMP-activated anion conductance in NMCM was permeable to both
Cl and ATP. Thus, although this electrodiffusional
anion pathway may be associated with the expression of an as yet
unidentified anion channel, these data are most consistent with the
expression of a functional CFTR, which was confirmed by
immunodetection. Expression of CFTR by Western blots of either neonatal
hearts or cultured NMCM were found to contain mature CFTR. Furthermore, the cAMP-activated currents were also consistent with a functional CFTR
phenotype, namely the activation of highly linear, time-independent currents in symmetrical Cl
, and a halide
perm-selectivity of Br
> Cl
> I
> gluconate. In addition, the cAMP-induced
Cl
currents were insensitive to DIDS but were
inhibited by DPC and glibenclamide, both known blockers of CFTR (20,
32). However, the blocking effect(s) of DPC and glibenclamide showed
little voltage dependence compared with previous reports on epithelial CFTR (20, 33). Nevertheless, the possibility exists for a stronger
competition between intracellular Cl
and DPC in this
preparation, since voltage dependence was actually observed in low
intracellular Cl
(Fig. 1B) and high
intracellular ATP (Fig. 8B), but not symmetrical Cl
(Fig. 8B).
Further evidence suggesting that the cAMP-activated
Cl currents are associated with a functional
expression of cardiac CFTR was provided by the blocking effect of
intracellular dialysis with an anti-CFTR antibody recently shown to
block human epithelial CFTR (27). Most consistent with the presence of
functional CFTR (4, 28), however, was the finding that cAMP stimulation
elicited DPC-inhibitable Cl
and ATP currents under
asymmetrical ATP/Cl
conditions. The
ATP/Cl
perm-selectivity ratio of 0.42 and 0.37 for
zATP =
2 and
4, respectively, was in
agreement with values previously reported for CFTR-expressing cells
(28) and purified CFTR (4). Furthermore, cAMP-activated
Cl
- and ATP-permeable currents have also been
observed in neonatal rat cardiac myocytes (43) known to express CFTR
(40).
Although the findings above are consistent with CFTR function in the
heart, cAMP stimulation has also been associated with the activation of
an osmotically activated Cl conductance likely
distinct from CFTR (24, 38). This was ruled out in the present study,
however, as cAMP-stimulated Cl
currents of NMCM were
blocked by the anti-CFTR antibody, while the osmotically activated
currents were not. These results suggest, therefore, that the
osmotically stimulated and cAMP-stimulated Cl
currents of NMCM are likely reflections of two distinct
electrodiffusional pathways.
To further confirm the role of CFTR in the cAMP-activated anion
conductance of NMCM, we took advantage of the recently developed cftr(/
) mouse (36) that lacks a functional gene
for CFTR. As expected from the lack of CFTR, cAMP stimulation in
symmetrical NaCl and MgCl2 solutions was without effect on
the cftr(
/
) NMCM. This was confirmed by the
inability to stimulate single-channel activity with ATP and PKA in
excised patches of NMCM. Likewise, cAMP stimulation did not elicit
whole cell currents under asymmetrical ATP/Cl
conditions in the cftr(
/
) NMCM. These results are
in agreement with the absence of immunodetectable CFTR in
cftr(
/
) NMCM (Fig. 7). However, the cAMP cocktail
was able to activate a Cl
- and ATP-permeable
conductance in heterozygous cftr(+/
) NMCM, suggesting
that one copy of the CFTR gene may be sufficient to elicit a response.
This may account for the lack of a cystic fibrosis phenotype in the
heterozygous littermates (36). In addition, the results of the
single-channel experiments confirmed that a normal gene, and thus a
functional CFTR, is necessary to observe the whole cell and
single-channel currents reported in this manuscript.
The role of CFTR in the cAMP-stimulated anion conductance in NMCM was
further verified with the single-channel data, which revealed that
addition of PKA and ATP to excised patches stimulated Cl-permeable channels only in cftr(+/+) and
cftr(+/
) NMCM but not in the
cftr(
/
) NMCM. These Cl
channels displayed single-channel conductances between 11 and 17 pS,
similar to those previously reported for cardiac CFTR (9, 22). Ehara
and Ishihara (9) reported an epinephrine-activated 13-pS
Cl
channel in ventricular myocytes. In a later
study, Nagel et al. (22) reported a PKA- and ATP-dependent
Cl
channel from guinea pig ventricular myocytes with
a single-channel conductance of 12 pS, which was identified as CFTR.
The single-channel conductance for the channels reported here,
therefore, are distinctly different from the single-channel conductance
of 28 pS reported for the outwardly rectifying Cl
channels (7) and 1.3 pS for the Ca2+-activated
Cl
channel (5). Thus the present data are in
agreement with the contention that CFTR is likely responsible for the
cAMP-stimulated Cl
conductance of NMCM.
Besides the data with the cftr(/
) mice and the
immunodetection of CFTR in cftr(+/+) NMCM, a strong functional
indication of the presence of CFTR is the cAMP-activated
electrodiffusional movement of ATP, thus far only elicited by CFTR (4,
28). Further evidence in this regard, therefore, was the cellular ATP release under CT-stimulated conditions, which is also consistent with
expression of CFTR, as previously reported (26). The CT-stimulated steady-state ATP release was ~18.0 × 10
15
mol/cell, in agreement with the ATP release of CFTR-expressing mouse
mammary carcinoma cells (26). The lack of a CT-stimulated release of
cellular ATP from cftr(
/
) NMCM also implicates
CFTR in the CT-stimulated ATP release in the wild-type NMCM.
The ability of CFTR to conduct ATP has been a subject of previous debate (34). However, a recent study from our laboratory has finally demonstrated that purified CFTR is indeed able to conduct ATP (4). Furthermore, other studies have demonstrated that other organic anions are able to permeate CFTR (19). A paradigm is emerging that suggests that the functional role of CFTR is indeed related to its function as an ATP-permeable pathway (4, 28).
The physiological role of CFTR in the human heart still remains largely
unclear. Despite recent reports indicating the presence of mRNA for
CFTR in human cardiac myocytes (41), functional assays have failed to
demonstrate a cAMP-stimulated Cl conductance (24).
However, children with cystic fibrosis present symptoms of cardiac
disease associated with acute heart failure (21). Furthermore, it has
also been suggested that the ventricular diastolic reserve is
diminished in cystic fibrosis patients (25). Thus the possibility
exists for a dysfunctional CFTR to be implicated in the onset of
cardiac disease in cystic fibrosis. Conversely, wild-type CFTR may
itself play an important but as yet undefined functional role in the
human heart.
Current dogma suggest that CFTR is a Cl-permeable
channel. However, Cl
channel activation has failed
to modify the maximal repolarization potential or the shape of the
action potential in preparations known to contain CFTR (11, 22). Thus
it could be argued that CFTR activation in the heart may not be largely
associated with a Cl
conductance, providing a likely
scenario for experimental conditions where functional CFTR has failed
to be detected. CFTR-mediated release of cellular ATP, which can act as
an autocoid by binding to purinergic receptors, may be part of a
regulatory mechanism that is necessary for regulating myocyte function
in the developing heart where autonomic regulation is not fully
developed (31). Although the source of extracellular ATP has not been
positively identified, cardiac extracellular ATP has been associated
with afterdepolarizations linked to the pericellular ATP concentration (37). Thus it is tempting to postulate that mechanisms such as CFTR
function may be implicated.
In conclusion, the data in this report are most consistent with the
presence of a functional CFTR in NMCM, where it is associated with the
activation of an anion-selective electrodiffusional pathway permeable
to both Cl and ATP. The presence of CFTR in neonatal
cardiac tissue, and the absence of a cAMP-activated conductance in the
cftr(
/
) NMCM, further indicate that CFTR is
responsible for cAMP-activated anion conductance in NMCM. Although
other explanations are still feasible, the fact that CFTR has not been
detected in cardiac tissue of adult mice (18) may suggest a
developmental role for cardiac CFTR. This functional role of CFTR
requires further investigation.
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ACKNOWLEDGEMENTS |
---|
We gratefully acknowledge Drs. Richard L. Sidman and Aizhong Li (New England Regional Primate Research Center, Department of Neurology, Harvard Medical School) for the kind gift of pregnant mice (C57BL/6, +/+, P100). In addition, we gratefully acknowledge Dr. Yanning Cui (New England Regional Primate Research Center, Department of Cardiovascular Medicine, Harvard Medical School) for her assistance in developing the technique for isolating and culturing NMCM, and Brandon M. Sullivan for superb technical advice in the development of the PCR technique.
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FOOTNOTES |
---|
This work was supported by a Grant-in-Aid from the American Heart Association and with funds contributed by the American Heart Association, Massachusetts Affiliate.
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. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: H. F. Cantiello, Renal Unit, Massachusetts General Hospital East, Bldg. 149 13th St., Charlestown, MA 02129 (E-mail: cantiello{at}helix.mgh.harvard.edu).
Received 1 March 1999; accepted in final form 8 September 1999.
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