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Vol. 273, Issue 5, C1650-C1656, November 1997
1 Department of Molecular and Integrative Physiology, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801; and 2 Department of Anatomy and Cell Biology, University of Cape Town Medical School, Cape Town, South Africa
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ABSTRACT |
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Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
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Weak channel blocker-induced noise analysis was used to determine the way in which the steroids aldosterone and corticosterone stimulated apical membrane Na+ entry into the cells of tissue-cultured A6 epithelia. Among groups of tissues grown on a variety of substrates, in a variety of growth media, and with cells at passages 73-112, the steroids stimulated both amiloride-sensitive and amiloride-insensitive Na+ transport as measured by short-circuit currents in chambers perfused with either growth medium or a Ringer solution. From baseline rates of blocker-sensitive short-circuit current between 2 and 7 µA/cm2, transport was stimulated about threefold in all groups of experiments. Single channel currents averaged near 0.3 pA (growth medium) and 0.5 pA (Ringer) and were decreased 6-20% from controls by steroid due to the expected decreases of fractional transcellular resistance. Irrespective of baseline transport rates, the steroids in all groups of tissues stimulated transport by increase of the density of blocker-sensitive epithelial Na+ channels (ENaCs). Channel open probability was the same in control and stimulated tissues, averaging ~0.3 in all groups of tissues. Accordingly, steroid-mediated increases of open channel density responsible for stimulation of Na+ transport are due to increases of the apical membrane pool of functional channels and not their open probability.
electrophysiology; sodium channels; tissue culture; cortical collecting ducts; kidney; noise analysis
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INTRODUCTION |
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Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
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WHEREAS STEROID HORMONES are known to stimulate Na+ transport in tight epithelia, the specific steps and mechanisms involved in steroid-mediated regulation of epithelial amiloride-sensitive apical membrane Na+ channels (ENaCs) are unknown. Both mineralocorticoids and glucocorticoids act over periods of hours to increase Na+ entry into the cells (19) via open channels (15). Because changes of open channel density may reflect changes of channel open probability (Po) and/or the density of functional channels (NT), the fundamental question has been whether regulation of transport occurs by way of Po and/or NT.
Using methods of blocker-induced noise analysis, Baxendale et al. (2) first reported that aldosterone and corticosterone stimulated transport by increase of NT in tissue-cultured A6 epithelia. In native tissues of rat renal cortical collecting ducts, patch clamp revealed that stimulation of transport by diet and aldosterone occurred by increase of NT (14). Similarly, A6 epithelia grown on rat tail collagen films respond to aldosterone by increase of NT (12). Because the ways in which cells regulate their channels may depend on the substrate on which A6 cells are grown (3, 11), we were led in scope of the experiments reported here to study a variety of passages of A6 cells (passages 73-112) originating in different laboratories on substrates other than collagen-coated Nuclepore membranes that were used in our original experiments (2) of steroid effects on Po and NT.
We report that aldosterone and corticosterone stimulate transport by increase of NT with no change of Po of amiloride-sensitive Na+ channels, regardless of substrate, serum, growth medium, or baseline expression of transport.
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MATERIALS AND METHODS |
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Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
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Cell culture. A6 cells from American Type Culture Collection (ATCC; Rockville, MD) at passage 69 were subcultured repeatedly in plastic flasks and used at passages 73-112. Three groups of experiments to be referred to as groups A, B, and C were done with differences not only in passage number but also the growth medium and the permeabilized substrates on which the tissues were grown. Cells in group A at passages 73-80 originated in Dr. R. L. Duncan's laboratory (Renal Division, Jewish Hospital, St. Louis, MO), where confluent tissues were grown on collagen-coated Nucleopore membranes (0.8-µm pore size; Worthington, Freehold, NJ), as described previously (9). Confluent monolayers were brought to Urbana for the experimental part of the studies that were carried out in 1985-1986. Cells in group B were purchased from ATCC at passage 69, subcultured, used at passage 75 with tissue growth on Millicell HA substrates (Millipore, Bedford, MA), and studied in 1994 in Urbana. Cells in group C were obtained as a gift to Dr. W. J. Els from Dr. W. Van Driessche, used at passages 107-112 with tissue growth on Millicell HA substrates, and studied in 1995 in Cape Town. The results of other groups of experiments were the same, with tissues grown on Transwell-Clear (Costar, Cambridge, MA) and Anocell (Whatman, Clifton, NJ) substrates and in a Leibovitz-Ham growth medium (16, 17), and will not be reported here.
Growth and perfusion media. The growth medium for group A experiments was a glutamine-, glucose-, and pyruvate-supplemented Dulbecco's modified Eagle's medium (D5648, Sigma Chemical, St. Louis, MO) diluted 15%. NaHCO3 (8 mM), penicillin (100 U/ml), streptomycin (100 µg/ml), insulin (5 mU/ml), and 10% fetal bovine serum (FBS) were added to this medium. Cells and tissues were maintained in a humidified incubator at 28°C with air containing 1.7% CO2. Ten days after seeding of the cells on the collagen-coated Nuclepore membranes, the tissues were fed serum- and insulin-free medium. The tissues were studied on days 14-26 in their control, steroid-depleted states of transport.
The growth medium for group B and C experiments was a Dulbecco's modified Eagle's medium (84-5022EC, GIBCO, Grand Island, NY) with N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES; 4 mM), penicillin (25 U/ml) and streptomycin (25 µg/ml) (17-719R, BioWhittaker, Walkersville, MD), and 10% FBS (Hyclone, Logan, UT). Cells and tissues were grown in the presence of humidified air containing 1% CO2 in an incubator at 28°C. Removal of the FBS before treatment with steroids was found to be unnecessary and was not done in these groups of experiments. We report the results of experiments in which control and steroid-treated tissues originated from the same lots of tissues. Before the day of an experiment, tissues were fed either growth medium (control) or the same medium containing exogenous steroid (0.27 µM aldosterone or corticosterone). Both control and steroid-treated tissues were studied at intervals between ~4 and 50 h postfeeding and handled in the same ways during transfer of the tissues from the inserts to continuous perfusion chambers designed for noise analysis (1). Group A tissues were perfused with a Ringer solution consisting of 100 mM NaCl, 2.4 mM KHCO3, 1.0 mM CaCl2, and 5 mM glucose. Group B and C tissues were perfused with growth medium minus the FBS and antibiotics, thereby ensuring that the tissues were being studied under essentially the same conditions under which they were grown.Electrical measurements.
The methods of study with blocker-induced noise analysis were identical
to those described in detail previously (6, 7, 10). After transfer to
the chambers, the tissues were short-circuited continuously for at
least 1 h to allow the macroscopic short-circuit currents
(Isc) to
stabilize. Thereafter, during periods of ~30 min, the apical
membranes of the cells were exposed in steps to increasing
concentrations (5-50 µM) of the weak
Na+ channel blocker
6-chloro-3,5-diaminopyrazine-2-carboxamide (CDPC; Aldrich Chemical,
Milwaukee, WI). Current noise at each blocker concentration
(B) was amplified, digitized, and Fourier transformed to
yield power density spectra from which the low-frequency plateaus (S0) and corner
frequencies
(fc)
of the induced Lorentzians were determined by nonlinear curve fitting
of the combined Lorentzians, "1/f" noise at the lower
frequencies and amplifier noise at the higher frequencies. Blocker on-
and off-rate coefficients
(kob and
kbo,
respectively) were calculated from the slopes and intercepts of
B 
2
fc plots,
respectively, yielding the blocker equilibrium constant
KB = kbo/kob.
IAmilsc, the single channel
currents at any B are
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(1) |
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(2) |
~1-2
s with mean open and closed times of several seconds). At the
equilibrium redistribution of channels, the fractional inhibition of
open channel density and thus Na+
entry into the cells is dependent on the
Po.
Extrapolation of the apparent values of
Po
at the various B values to zero B circumvents the
autoregulatory increases of
NBo that lead to
underestimates of the
Po at any
B (10; see RESULTS). As
will be evident in
RESULTS, the
Po
values of blocker-sensitive channels estimated by noise analysis as
indicated above are virtually the same as those of the long open time
channels measured by patch clamp of A6 epithelia, underscoring the
validity of the assumption that blocker interacts only with the open
state of the channels in A6 epithelia as in those of frog skin.
The channel densities determined this way are a measure of the pool of
apical membrane channels directly involved in
Na+ transport. Hence,
INa = iNaPoNT,
where the functional
NT = No/Po. Values are means ± SE.
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RESULTS |
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Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
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Blocker-sensitive and blocker-insensitive Isc. Compared with previous published reports in studies of frog skin (6, 7, 10), A6 epithelia behaved in identical ways in response to CDPC inhibition of apical membrane Na+ channels, except the control baseline rates of transport were generally considerably less than those in native tissues (e.g., frog skin, toad urinary bladder, cortical collecting duct, and others). As indicated in Fig. 1, the strip chart recordings of the changes of Isc in response to staircase increases of blocker concentration showed the typical scalloped appearance, indicative of autoregulatory increases of channel density (1, 10), and the typical overshoot of the Isc after complete washout of CDPC from the apical solution. From peak values, the currents relaxed toward the original control Isc within ~10 min. Addition of 100 µM amiloride to the apical solution caused marked inhibition of the Isc, but not to zero, leaving an IAmilsc. Removal of all Na+ from the Ringer solution perfusing the tissues (group A) caused the IAmilsc to fall to zero, indicating that the IAmilsc was due to highly selective Na+-conductive but blocker-insensitive channels.
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Single channel currents and open channel densities.
Despite the low baseline values of transport (compared with native
tissues like frog skin and toad urinary bladder), noise analysis was
possible at
Isc
somewhat less than 1 µA/cm2 at
B 
5 µM CDPC. As indicated in Fig.
2, Lorentzians were easily resolved at
5-50 µM CDPC. For all groups of experiments and as indicated for
group A experiments summarized in Fig.
3, the
fc varied linearly with B in both control and steroid-treated
tissues. S0
and IBNa of control and
steroid-treated tissues changed in accordance with a kinetic scheme of
closed 
open blocked states, where the blocker interacts only
with the open state of the channel. As indicated in Fig. 3 for
group A tissues and summarized in
Table 1 for all groups, the
iNa
were less in steroid-treated tissues than in controls due at least in
part to depolarization of apical membrane voltage in their stimulated
states of transport (3, 7).
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Blocker on- and off-rates. The kob and kbo measured in A6 epithelia, summarized in Table 2, were virtually the same as those reported for Na+ channels in frog skin. Accordingly, the blocker site appears to be conserved between these tissues. Both the access time to the blocker site and the residency time at the site were essentially the same among all passages of cells and conditions of growth, yielding KB that averaged ~30 µM. It was apparent that the channels recruited by steroid possessed virtually the same kinetic interactions with CDPC.
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Po and NT. It is evident from the data summarized in Fig. 5 that steroid stimulation of transport does not occur by increase of Po. Po averaged between ~0.25 and 0.35 in all groups of tissues and was not changed by steroid treatment of the tissues. The mean values of Po measured here by noise analysis are virtually the same as those of the long open time channel described by Kemendy et al. (12) in their patch-clamp studies of A6 epithelia.
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DISCUSSION |
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Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
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Regardless of baseline rates of Na+ transport, age and origin of the cells, substrates on which the cells were grown, growth media, and serum, our results demonstrate that the steroid hormones aldosterone and corticosterone increase Na+ transport by increase of the density of functional channels at the apical membranes of the cells. The channels are the same, as judged by the similarity in the kinetics of blocker interaction of the channels and by the magnitudes of the single channel currents that are in the same range as those measured by patch clamp and those expected for single channel conductance near 5 pS (3). In comparison with frog skin, the channels appear to be identical, possessing extremely high selectivity for Na+ where the predominant population of channels is amiloride inhibitable. Because the Po is not changed by aldosterone or corticosterone, stimulation of transport must be due to increase in the apical membrane pool size of functional channels involved in Na+ transport. It is widely appreciated that channels may be recruited from pools of nonfunctional channels present in the apical membranes and/or trafficked to the apical membrane from the cytosol. Our experiments do not address this issue or the mechanism whereby steroids increase NT. Using identical methods of blocker-induced noise analysis, Granitzer et al. (8) reported that the glucocorticoid dexamethasone stimulates NT in A6 epithelia with no change of Po. Pácha et al. (14) reported that Po was not influenced by mineralocorticoid status in the rat cortical collecting duct; in contrast, Kemendy et al. (12) reported that aldosterone shifted channels from a short to a long open time state in A6 cells. Accordingly, regardless of the origin of the channels, Na+ transport must be carried out principally by blocker-sensitive channels with long open and closed times and by channels in which Po is the same at various states of mineralocorticoid and glucocorticoid status.
Blocker-sensitive and blocker-insensitive
Isc.
Since discovery of amiloride as a potent inhibitor of epithelial
Na+ transport, it has been widely
acknowledged that this diuretic inhibits most, but not all, of the
Isc in target
tissues like frog skins and toad urinary bladders. The same appears to
be true for A6 epithelia in steroid-depleted and steroid-stimulated A6 epithelia. Regardless of the permeabilized substrate on which the cells
are grown (11) and as indicated in Table 1, the amiloride- or
blocker-insensitive currents 1)
represent a rather small fraction of the macroscopic rates of transport
into the cells as measured in absolute and relative terms and
2) are stimulated by steroid hormone. Because the IAmilsc
decreases to zero after removal of all
Na+ from the apical solution, the
apical membrane must contain channels that are conductive to
Na+ but that are not blocked by
amiloride at concentrations 100 µM that far exceed by several
orders of magnitude the KAmilB of blocker-sensitive channels. Accordingly, apical membranes must possess blocker-sensitive and blocker-insensitive pools of channels and
corresponding blocker-sensitive and blocker-insensitive
Isc. Because
blocker-induced noise analysis measures only those channels that are
blocker sensitive and because blocker-insensitive
Na+ currents represent a very
small fraction of Na+ transport,
increases of transport must be attributable to increases of
blocker-sensitive
NT.
Nevertheless, blocker-insensitive currents are also increased by
steroids but represent a comparatively small fraction of
Na+ transport in the presence and
absence of exogenous steroids. The gating kinetics and single channel
conductance of blocker-insensitive Na+ channels are unknown.
o ~ 40 ms) coexisted with
mean long open time channels (o ~ 1,600 ms) in the same patches. It is unknown whether the short open
time channels observed by patch clamp are blocker sensitive or
insensitive. Thus the role and function of short open time channels
remain unknown but could perhaps be related to blocker-insensitive Na+ channels. If the densities of
short and long open time channels are in the same range, then the
contribution of short open time channels to the macroscopic rates of
Na+ transport would be in the
range of a few percent of the
Isc. With
identical channel densities and closed times and with
o of 40 and 1,600 ms, the
contribution of short open time channels to the
Isc is 2.5%.
Origin of channels. It is of particular interest to know the origin of the long open time channels, the density of which is increased by steroids. The hypothesis by Kemendy et al. (12) that steroids change open probability of preexisting short open time channels into long open time channels by an all-or-none mechanism rests on critical observations. First, short open time channels must exist before exposure of the tissues to steroids. Second, assuming that the density of short open time channels is constant and that steroids stimulate transport by an all-or-none increase of the mean open time of these channels, decreases of short open time channels must be accompanied by identical increases of long open time channels if this is the only source of channels.
It is appreciated that interpretation of patch-clamp data is exacerbated, especially for the particular case where channel open times are small compared with their closed times (4), as is the case for the short open time channels. With 40-ms mean open time and 3,000-ms mean closed time of the short open time channels, there would, at a 95% level of confidence, have to be at least 13 channels in a patch to observe a double opening of this channel within 20 min of continuous recording (4, 12). The probability of observing multiple openings of these channels is even more remote, so estimates of short open time channel number in a patch are practically impossible. Accordingly, it would be impossible practically to know whether steroids cause changes of short open time channel densities and, hence, whether long open time channels originated from short open time channels as has been suggested (12). The source of the apical pool of functional channels is unknown and remains a topic of particular interest. Our own experiments reported here shed no light on this issue. Our experiments do not rule out sources of nonfunctional channels within the apical membrane or sources originating from channel-containing intracellular vesicles. Aldosterone is known to induce a variety of proteins, among which may be channel subunits and other proteins involved in sorting and trafficking the channels to the apical membranes. Subunits are expressed in steroid-depleted tissues, and aldosterone does not change the levels of subunit RNA in the same way in all tissues, if at all (13). cDNA-injected oocytes express long open time channels (18) with no reports of short open time channels like those observed in A6 (12). Because apical membrane capacitance is increased by aldosterone in tissues treated overnight (13a) and in a time-dependent way during exposure to aldosterone for 6 h (11a), it is possible that steroids stimulate transport by trafficking of channels to the plasma membrane. Accordingly, steroids stimulate Na+ transport by increase of apical membrane NT with no change of Po. The question of origin of the channels remains open.| |
ACKNOWLEDGEMENTS |
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We thank A. L. Helman for excellence in maintaining our tissue culture facility (Urbana, IL), the care and feeding of the cells, and assistance in the preparation of the manuscript.
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FOOTNOTES |
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This study was supported by National Institutes of Health Grants DK-16663 and DK-30824 (to S. I. Helman) and RR-05491 (to R. L. Duncan) and grants from the South African Medical Research Council and the National Kidney Foundation of South Africa (to W. J. Els).
L. M. Baxendale-Cox was an American Heart Association (Illinois Affiliate) Fellow. X. Liu is a doctoral student in the Dept. of Molecular and Integrative Physiology (Urbana). K. Baldwin is a master's degree student in the Dept. of Anatomy and Cell Biology (Cape Town, South Africa).
Present addresses: L. M. Baxendale-Cox, School of Nursing, Johns Hopkins University, Baltimore, MD 21205; R. L. Duncan, Dept. of Orthopaedic Surgery, Indiana University School of Medicine, Indianapolis, IN 46202.
Address for reprint requests: S. I. Helman, Dept. of Molecular and Integrative Physiology, 524 Burrill Hall, 407 South Goodwin Ave., University of Illinois at Urbana-Champaign, Urbana, IL 61801.
Received 21 February 1997; accepted in final form 8 July 1997.
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REFERENCES |
|---|
|
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
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1.
Abramcheck, F. J.,
W. Van Driessche,
and
S. I. Helman.
Autoregulation of apical membrane Na+ permeability of tight epithelia. Noise analysis with amiloride and CGS 4270.
J. Gen. Physiol.
85:
555-582,
1985
2.
Baxendale, L. M.,
R. L. Duncan,
and
S. I. Helman.
Aldosterone increases apical membrane Na+ channel density in A6 epithelia (Abstract).
Federation Proc.
46:
495,
1987.
3.
Blazer-Yost, B. L.,
and
S. I. Helman.
The amiloride-sensitive epithelial Na+ channel: binding sites and channel densities.
Am. J. Physiol.
272 (Cell Physiol. 41):
C761-C769,
1997
4.
Colquhoun, D.,
and
A. G. Hawkes.
The principles of the stochastic interpretation of ion-channel mechanisms.
In: Single-Channel Recording, edited by B. Sakmann,
and E. Neher. New York: Plenum, 1983, p. 135-175.
5.
Eaton, D. C.,
and
Y. Marunaka.
Ion channel fluctuations: "noise" and single-channel measurements.
In: Channels and Noise in Epithelial Tissues (Current Topics in Membranes and Transport), edited by S. I. Helman,
and W. Van Driessche. New York: Academic, 1990, vol. 37, p. 61-113.
6.
Els, W. J.,
and
S. I. Helman.
Activation of epithelial Na channels by hormonal and autoregulatory mechanisms of action.
J. Gen. Physiol.
98:
1197-1220,
1991
7.
Els, W. J.,
and
S. I. Helman.
Dual role of prostaglandins (PGE2) in regulation of channel density and open probability of epithelial Na+ channels in frog skin (R. pipiens).
J. Membr. Biol.
155:
75-87,
1997[Medline].
8.
Granitzer, M.,
I. Mountian,
and
W. Van Driessche.
Effect of dexamethasone on sodium channel block and densities in A6 cells.
Pflügers Arch.
430:
493-500,
1995[Medline].
9.
Grogan, W. M.,
M. L. Fidelman,
D. E. Newton,
R. L. Duncan,
and
C. O. Watlington.
A corticosterone metabolite produced by A6 (toad kidney) cells in culture: identification and effects on Na+ transport.
Endocrinology
116:
1189-1194,
1985
10.
Helman, S. I.,
and
L. M. Baxendale.
Blocker-related changes of channel density. Analysis of a three-state model for apical Na channels of frog skin.
J. Gen. Physiol.
95:
647-678,
1990
11.
Helman, S. I.,
and
X. Liu.
Substrate-dependent expression of Na+ transport and shunt conductance in A6 epithelia.
Am. J. Physiol.
273 (Cell Physiol. 42):
C434-C441,
1997
11a.
Helman, S. I.,
X. Liu,
and
B. Blazer-Yost.
Early response of A6 epithelia to aldosterone is mediated by vesicle trafficking of apical Na+ channels and not open probability (Abstract).
FASEB J.
10:
A78,
1996.
12.
Kemendy, A. E.,
T. R. Kleyman,
and
D. C. Eaton.
Aldosterone alters the open probability of amiloride-blockable sodium channels in A6 epithelia.
Am. J. Physiol.
263 (Cell Physiol. 32):
C825-C837,
1992
13.
Lingueglia, E.,
N. Voilley,
M. Lazdunski,
and
P. Barbry.
Molecular biology of the amiloride-sensitive epithelial Na+ channel.
Exp. Physiol.
81:
483-492,
1996[Abstract].
13a.
Liu, X.,
and
S. I. Helman.
Aldosterone increases apical membrane capacitance of A6 epithelia (Abstract).
FASEB J.
9:
A64,
1995.
14.
Pácha, J.,
G. Frindt,
L. Antonian,
R. B. Silver,
and
L. G. Palmer.
Regulation of Na channels of the rat cortical collecting tubule by aldosterone.
J. Gen. Physiol.
102:
25-42,
1993
15.
Palmer, L. G.,
J. H.-Y. Li,
B. Lindemann,
and
I. S. Edelman.
Aldosterone control of the density of sodium channels in the toad urinary bladder.
J. Membr. Biol.
64:
91-102,
1982[Medline].
16.
Paunescu, T. G.,
and
S. I. Helman.
Dual role of prostaglandin E2 in regulation of Na+ transport in A6 epithelia (Abstract).
Biophys. J.
72:
A230,
1997.
17.
Paunescu, T. G.,
X. Liu,
and
S. I. Helman.
Nonhormonal regulation of apical membrane sodium transport in A6 epithelia (Abstract).
FASEB J.
11:
A8,
1997.
18.
Puoti, A.,
A. May,
C. M. Canessa,
J.-D. Horisberger,
L. Schild,
and
B. C. Rossier.
The highly selective low-conductance epithelial Na channel of Xenopus laevis A6 kidney cells.
Am. J. Physiol.
269 (Cell Physiol. 38):
C188-C197,
1995
19.
Watlington, C. O.,
F. M. Perkins,
P. J. Munson,
and
J. S. Handler.
Aldosterone and corticosterone binding and effects of Na+ transport in cultured kidney cells.
Am. J. Physiol.
242 (Renal Fluid Electrolyte Physiol. 11):
F610-F619,
1982.
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),
and noise at higher frequencies
(S2f
),
originating at input stage of voltage amplifier. Spectra at intermediate concentrations are not shown.
