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secretion but not
Na+ reabsorption in A6
cells
Departments of 1 Physiology and Biophysics, 2 Cell Biology, and 3 Medicine, University of Alabama at Birmingham, Birmingham, Alabama 35294
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ABSTRACT |
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Top
Abstract
Introduction
Methods
Results
Discussion
References
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The effects of microtubule disruption on arginine
vasotocin (AVT)-stimulated Na+ and
Cl
transport were studied
in A6 cells by measuring short-circuit currents
(Isc) across
cell layers grown in tissue culture on permeable supports. Microtubule
disruption inhibited an AVT-stimulated secretory Cl current but did not
prevent activation of amiloride-sensitive Na+ transport. This AVT-stimulated
secretory Cl current was
significantly inhibited by glibenclamide, an inhibitor of
the cystic fibrosis transmembrane conductance regulator (CFTR). Reverse
transcription of RNA isolated from A6 cells followed by polymerase
chain reaction (PCR) using primers designed to amplify a portion of the
R-domain of CFTR cloned from Xenopus
laevis skin and immunocytochemistry demonstrated the
presence of CFTR in A6 cells and an apparent recruitment of cytoplasmic
CFTR to the apical cell surface after AVT stimulation. In contrast,
indirect immunofluorescent labeling of
Na+ channels using a polyclonal
antibody raised against a biochemically isolated
Na+ channel complex from bovine
renal medulla labeled the apical plasma membrane but failed to
demonstrate intracellular labeling of
Na+ channels (except in
subconfluent cells) or recruitment of
Na+ channels to the apical
membrane region after AVT stimulation.
trafficking; cystic fibrosis transmembrane conductance regulator; cytoskeleton; amiloride-sensitive sodium channel; sodium transport; chloride transport
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INTRODUCTION |
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Top
Abstract
Introduction
Methods
Results
Discussion
References
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A6 CELLS, ORIGINALLY derived as a continuous
cell line from Xenopus kidney, have
been used extensively to study amiloride-sensitive Na+ transport. Filter-grown A6
cells display characteristics similar to the mammalian cortical
collecting duct: a tight epithelium with amiloride-sensitive,
electrogenic Na+ transport that is
stimulated by both arginine vasotocin (AVT) and aldosterone. In
addition to Na+ channels, A6 cells
also have Cl channels in
their apical membranes (19) that may be involved in an AVT-stimulated
Cl conductance (8, 32, 34).
AVT, as well as its mammalian homolog arginine vasopressin (AVP), have
been shown to stimulate water and/or
Na+ transport in a variety of
epithelia including rat collecting duct segments (9) and primary
cultures of renal collecting tubules (5), as well as amphibian analogs
such as A6 cells (2). In some epithelia, AVT- or AVP-stimulated
increases in water permeability are dependent on the insertion of water
channels into the cell membrane. The "shuttling" of transporters
into and out of cell membranes as a means of regulating the transport
of a variety of substances has been proposed for several different tissues: the GLUT-4 glucose transporter in skeletal muscle (14), the
cystic fibrosis transmembrane conductance regulator (CFTR) anion
channel in Cl-secreting
epithelia such as the distal colon (31), and the aquaporin-2 (AQP-2)
water channel in the renal collecting duct (18). However, there is
debate as to whether shuttling of
Na+ channels into the apical
membrane of A6 and other epithelial cells is responsible for the
increase in Na+ transport observed
after stimulation with AVT.
Two mechanisms have been advanced to explain the increased Na+ transport upon stimulation with AVT: 1) insertion of Na+ channels into the apical membrane (16) or 2) phosphorylation of Na+ channels already residing in the apical membrane (15, 21). Kleyman et al. (16) have presented evidence for the former by using radioiodination of cell surface proteins and subsequent immunoprecipitation of Na+ channels with an anti-idiotypic antibody to demonstrate an increase in apical expression of Na+ channels after AVT-stimulation. Additional evidence for AVT-stimulated insertion of Na+ channels was presented by Marunaka and Eaton (20), who showed that AVT and cAMP increase the number of Na+ channels per patch in A6 cells, presumably independent of changes in open probability. On the other hand, evidence for activation of resident channels has come from the work of Benos and colleagues (26), who have shown that AVT treatment of cultured A6 cells results in phosphorylation of a 300-kDa polypeptide associated with a biochemically purified renal Na+ channel complex (26) and, in planar lipid bilayer experiments, that protein kinase A (PKA)-mediated phosphorylation of biochemically purified Na+ channels from bovine renal medulla changes single channel characteristics enabling greater movement of Na+ through the channels (15, 21).
Additional evidence suggesting that antidiuretic hormone (ADH; refers to either AVP or AVT) stimulates Na+ transport via an activation of Na+ channels already present in the apical membrane comes from previous studies using toad bladder (17, 22, 28, 29). These experiments have clearly demonstrated that colchicine (29) (a disrupter of microtubule structure), cytochalasin B (22) (a disrupter of filamentous actin structures), trifluoperazine (17), and methohexital sodium (28) dissociate the effects of ADH on water transport from its effects on Na+ transport; each of these agents inhibits ADH-stimulated water transport but was without effect on ADH-stimulated short-circuit current (Isc). Stetson et al. (28) and Palmer and Lorenzen (22) have also reported that ADH-stimulated increases in capacitance (as a measure of membrane surface area) are correlated with increased water transport but not Na+ transport; furthermore, the ADH-stimulated increases in capacitance and water transport in the toad bladder can be inhibited by methohexital, colchicine, and cytochalasin B, without effect on ADH-stimulated Na+ transport. These observations suggest either that ADH-stimulated increases in Na+ transport are not due to a vesicle fusion or insertion of "new" Na+ channels or that ADH-stimulated trafficking of Na+ channels occurs through a pathway different from trafficking of water channels.
Various cytoskeletal components, including actin filaments and
microtubules, have been implicated in regulating a number of transport
processes (6, 7, 23) and also appear to be involved in various aspects
of vesicular transport including endo- and exocytosis (33). The aim of
this study was to address what role the cytoskeleton, particularly the
microtubules, plays in AVT-stimulated Na+ and
Cl transport in A6 cells.
We present here evidence that microtubule disruption inhibits an
AVT-stimulated Cl but not
Na+ transport.
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METHODS |
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Top
Abstract
Introduction
Methods
Results
Discussion
References
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Cell culture. A62F3 cells, a subclone of A6 cells (a gift from B. Rossier, Lausanne, Switzerland), were cultured in T-75 culture flasks at 26°C in a humidified incubator in the presence of 1% CO2. The culture medium was a specially formulated amphibian variant of Dulbecco's modified Eagle's medium (GIBCO-BRL; formula 84-5022EL) containing 74 mM NaCl and 8 mM HCO3, 195 mosmol/kgH2O. To this was added 2% penicillin and streptomycin and 10% calf bovine serum (CBS). Cells were fed twice weekly and split upon reaching confluence. Cells were seeded on 24-mm Cyclopore (Fisher Scientific, Atlanta, GA) inserts at an approximate density of 1.6 ×106 cells/well, placed in 6-well companion tissue culture plates, and fed twice weekly with medium containing 10% CBS until a transepithelial potential (VT) of at least 20 mV developed. Inserts having a potential of 20 mV (apical side negative) were considered viable for experiments; however, inserts typically had potentials of 30-40 mV (usually 11 days after seeding). Cells older than 16 days were discarded.
Cytoskeletal disruption. Drugs or
vehicle from stock vials [10 mM colchicine and 
-lumicolchicine
in dimethyl sulfoxide (DMSO), 1 mg/ml nocodazole in DMSO, and 1 mM
cytochalasin E in DMSO] were added to A6 medium (without CBS)
to a final concentration of 100 µM colchicine or
-lumicolchicine, 33 µM nocodazole, or 500 nM cytochalasin E. Culture medium was removed from cells both apically and basolaterally,
replaced with medium containing drugs (or the appropriate vehicle in
controls), and incubated accordingly: colchicine, -lumicolchicine,
and nocodazole for 30 min at 4°C followed by 3 h at 26°C; and
cytochalasin E, 1 h at 26°C. Following incubation, the
bottoms of the inserts were removed with a scalpel and either processed
for immunocytochemistry or mounted in Ussing-type chambers for
Isc measurements.
Cell fixation and immunocytochemistry for
actin. Inserts were briefly rinsed with
phosphate-buffered saline (PBS), then fixed and permeabilized with
high-performance liquid chromatography (HPLC)-grade absolute acetone
for 10 min at 
20°C. Inserts were rinsed with PBS (137 mM
NaCl, 2.7 mM KCl, 1.5 mM
KH2PO4,
and 8.0 mM
Na2HPO4,
pH 7.4), then postfixed with 3% formaldehyde in PBS for 15 min, in the
dark, at room temperature. Monolayers were then rinsed with PBS and
blocked with 1% bovine serum albumin (BSA) in PBS for 30 min. Next,
inserts were incubated with rhodamine-phalloidin (Boehringer-Mannheim,
Indianapolis, IN), diluted 1:20 in 1% BSA/PBS for 1 h at 37°C,
rinsed, and mounted cell-side-up on glass slides. The mounting medium
consisted of 0.1% p-phenylenediamine
in 1:9 PBS/glycerol.
Cell fixation and immunocytochemistry for
tubulin. Inserts were briefly rinsed with PBS and
permeabilized with 1% Triton in PEM [0.1 M
piperazine-N,N'-bis(2-ethanesulfonic
acid), 1 mM ethylene glycol-bis(-aminoethyl
ether)-N,N,N',N'-tetraacetic
acid, 1 mM MgCl2, pH 6.9]
for 2.5 min. Inserts were then rinsed four times with PEM and fixed
with 3% formaldehyde for 45 min in the dark at room temperature. The
inserts were again rinsed with four changes of PEM and postfixed in
HPLC-grade absolute acetone for 30 min at 20°C. Inserts were
then rinsed with four additional changes of PEM and prepared for
immunocytochemistry. The filters were blocked with 1% BSA in PBS for
30 min and then incubated with Tu27B (diluted 1:20 in 1% BSA), a mouse
monoclonal anti-tubulin antibody (from Lester I. Binder, Northwestern
University, Chicago, IL) for 1 h at 37°C. After washing, the
inserts were serially incubated with a secondary antibody, fluorescein
isothiocyanate (FITC)-conjugated goat anti-mouse
immunoglobulin G (IgG), and with a tertiary antibody, swine anti-goat
IgG-FITC (each diluted 1:25 in 1% BSA/PBS) for 1 h each,
at 37°C. Inserts were then stained with Hoechst 33258 dye (20 µg/ml in PBS), to localize cell nuclei, and mounted on slides,
cell-side-up. The mounting medium was the same as for the actin
immunocytochemistry.
Cell fixation and immunocytochemistry for
Na+
channels and CFTR.
Paired control and experimental cell culture inserts were examined in
parallel by measuring the
VT. After any
pretreatment with agents such as nocodazole or colchicine, AVT was
added, and when
VT had reached an
apparent maximum (15 to 20 min after AVT addition), the filter bottoms
were cut out of the plastic inserts and were briefly rinsed with PBS
(220 mosmol/l) and then fixed and permeabilized with 3:1
methanol/acetic acid (vol/vol) for 30 min at 20°C. Inserts
were washed with four changes of PBS and postfixed with 3%
formaldehyde in PBS for 15 min at room temperature in the dark. After
again washing with four changes of PBS, the filters were blocked with
1% BSA in PBS for 15 min. For Na+
channel localization, filters were then incubated for 1 h at 37°C
with a rabbit polyclonal antibody made against
Na+ channel purified from bovine
medulla (30), diluted 1:20 in 1% BSA in PBS to a final concentration
of 80 µg/ml. Preimmune controls were prepared in the same manner. For
CFTR localization, filters were incubated for 1 h at 37°C with a
mouse monoclonal antibody against the regulatory (R)-domain of CFTR
(Genzyme, Boston, MA) (10). After being washed, the filters were then
serially incubated with either FITC-conjugated secondary goat
anti-rabbit IgG (Boehringer-Mannheim) or FITC-conjugated goat
anti-mouse IgG diluted 1:20 in 1% BSA in PBS for
Na+ channel or CFTR
immunofluorescence, respectively, followed by FITC-conjugated swine
anti-goat tertiary antibody (Boehringer-Mannheim) diluted
1:20 in 1% BSA/PBS, each for 1 h at 37°C. Nuclei were then stained
with Hoechst 33258 (see above), and samples were mounted in cross
section by filter folding (30). The same mounting medium was used for
actin and tubulin immunocytochemistry. Quantification of
immunofluorescence was performed as previously described (12).
Ringer"
solution containing (in mM) 95 sodium gluconate, 5 NaCl, 1 calcium
gluconate, 0.5 magnesium gluconate, 2 K2HPO4,
10 HEPES, and 5 glucose was used to identify the
Cl-dependent components of
the AVT-stimulated
Isc.
At least two monolayers from the same plating and incubated under the
same conditions were used in each experiment, with one monolayer
serving as a control and one or two monolayers for experimental maneuvers. Statistical comparisons between control and experimental monolayers were based on a two-tailed
t-test for significance. When a
maneuver such as AVT addition was performed, the significance of the
effect was assessed based on a t-test
of the mean paired difference for all preparations undergoing the same
treatment. Significance was assumed at
P < 0.05.
RNA isolation and RT-PCR. Total RNA
was isolated from A6 cells using TRIzol (Life Technologies,
Gaithersburg, MD) according the manufacturer's recommendations. The
RNA samples were treated with ribonuclease-free deoxyribonuclease I
(Life Technologies) and then reverse transcribed with SuperScript II
(Life Technologies) using random hexamers, 2 µl per 100-µl reaction
(Life Technologies). Aliquots (10 µl) of the reverse transcription
(RT) reaction were removed and were run without SuperScript II and used
as a control for genomic DNA contamination in subsequent polymerase
chain reaction (PCR) amplifications. Because A6 cells derive from
Xenopus kidney, RNA was extracted from
a ventral section of Xenopus laevis
skin to serve as a positive control.
The cDNA from A6 cells was probed for the presence of CFTR by PCR using
primers designed to amplify a portion of the R-domain of the cloned
sequence from Xenopus skin (11) (Note:
at the time our studies were done, CFTR from A6 cells had not yet been sequenced): forward (CFTR.A6.S), 5' GAA GGT GGA ATT ACA TTA AGT GGA G 3'; reverse (CFTR.A6.AS), 5' CAG GTC AAA TGA TGA GTT
GGA G 3'.
PCR reactions were performed in 50-µl total reaction volumes
containing 50 mM KCl, 20 mM tris(hydroxymethyl)aminomethane
hydrochloride (pH 8.4), 2.5 mM
MgCl2, 0.1 mg/ml BSA, 0.1 mM of
each deoxynucleoside 5'-triphosphate, 25 pmol of each primer, and
2 U of Taq polymerase (Promega,
Madison, WI). The PCR amplifications were performed in a Perkin-Elmer
DNA Thermal Cycler with the following cycler protocol: 94°C initial
melt for 3 min followed by 94°C for 1 min, 56°C for 1 min,
72°C for 1 min, (30 cycles), with a 7-min extension at 72°C at
the end of the cycling. The PCR reaction products were resolved on 2%
agarose gels.
PCR amplification products were TA cloned per the manufacturer's
instructions (Invitrogen, Carlsbad, CA). The clone was then sequenced
in both directions using T7 and M13 universal sequencing primers and
dye-termination reactions at the University of Alabama DNA Sequencing
Core Facility (Dr. S. Hollingshead, Director). The degree of sequence
homology to known sequences from A6 and Xenopus was analyzed using the
Wisconsin Sequence Analysis Package (version 8; Genetics Computer
Group, Madison, WI).
Sources of biochemicals and stock
solutions. In addition to the sources of biochemicals
mentioned in the preceding text. The following biochemicals were
obtained from Sigma (St. Louis, MO): amiloride, AVT,
-lumicolchicine, BSA, colchicine, cytochalasin E,
4,4'-diisothiocyanostilbene-2,2'-disulfonic acid (DIDS),
glibenclamide, Hoechst 33258 dye, nocodazole, and
p-phenylenediamine. All other chemicals were obtained in purest available grade from Fisher. A stock
solution of 10 µM AVT was made with PBS (pH 6.8), which was added to
the bathing solution for a final concentration of 0.1 µM. Amiloride
was added from a 10 mM stock solution in water. Glibenclamide was
disolved in DMSO to give a 30 mM stock solution. A 100 mM DIDS stock
solution was made in water. Whenever a vehicle other than water or PBS
was used in a stock solution, control cells received an equivalent
amount of the vehicle alone.
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RESULTS |
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Top
Abstract
Introduction
Methods
Results
Discussion
References
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The effect of nocodazole and colchicine treatment on microtubule structure was verified with immunocytochemistry. Both nocodazole and colchicine treatment disrupted microtubule structure, with nocodazole appearing to be more effective than colchicine, at least at the concentrations used (Fig. 1). The effect of 500 nM cytochalasin E on actin was also visualized with immunocytochemistry, but only the basolateral actin comprising the stress fibers was found to be affected. The cortical actin in the apical and lateral domains appeared largely intact after a 1-h incubation with 500 nM cytochalasin E (data not shown). In accordance with previous observations (23), incubation of A6 cells for 1 h with micromolar concentrations of cytochalasin E structurally disrupts the cells, precluding their use in transport studies.
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Figure 2 is a representative experiment showing the effects of microtubule disruption by nocodazole on the Isc and G responses to 0.1 µM AVT. In both the control and nocodazole-treated cells, addition of AVT resulted in an increase in Isc and G (indicated by lengthening of the voltage-induced deflections in Isc). When 10 µM amiloride was added after 90-min exposure to AVT, Isc was reduced to zero in both control and nocodazole-treated cells. In this experiment as in all experiments in this series, both the basal and AVT-stimulated Isc were decreased by nocodazole. This inhibitory effect of nocodazole (and also of colchicine) is considered below, but we focus first on the time course of the Isc response to AVT. The most obvious difference in the pattern of the response between the control and nocodazole-treated cells in Fig. 2 occurs in the first 30 min after AVT addition. Figure 3 compares the time courses for Isc in Fig. 2 and those of two other experiments in this series. In the three control preparations shown in Fig. 3, within 15-25 min AVT produced an initial increase in Isc that was followed by a decline to a slightly reduced level. In contrast, AVT produced a slower and monotonic rise in Isc in the four nocodazole-treated preparations. The same difference in the time course of Isc is seen when one compares the AVT responses of three control preparations with those of five colchicine-treated preparations (Fig. 4). Again, the control cells exhibited a biphasic Isc response to AVT with an early peak, whereas the colchicine-treated cells showed only a slower monotonic increase in Isc with AVT.
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In the series of experiments presented in Fig. 3, the control preparations had a higher basal Isc than the nocodazole-treated preparations (15.3 ± 2.7 vs. 5.1 ± 0.4 µA/cm2, respectively). AVT increased the Isc to a steady-state value of 38.2 ± 4.1 in control vs. 19.7 ± 1.9 µA/cm2 in nocodazole-treated cells. Thus, although the absolute increment in Isc produce by AVT was less in nocodazole-treated monolayers than control monolayers, the relative increment (the ratio of "steady-state" or plateau Isc to basal Isc) was actually greater for nocodazole-treated monolayers than control monolayers (3.9-fold vs. 2.5-fold, respectively). Similarly, in the experiments shown in Fig. 4, control preparations had a basal Isc of 8.7 ± 4.0 vs. 3.2 ± 0.8 µA/cm2 in colchicine-treated monolayers. AVT increased Isc to 16.7 ± 1.6 and 6.3 ± 1.7 µA/cm2 in control and colchicine-treated cells, respectively. Again, the absolute magnitude of the AVT-stimulated Isc was less in monolayers with microtubule disruption compared with controls; however, the relative increases in Isc were the same for both control (1.9-fold) and colchicine-treated (2.0-fold) monolayers. AVT also significantly (P < 0.01) increased G in the six control preparations in Figs. 3 and 4 from 0.58 ± 0.09 to 0.93 ± 0.05 mS/cm2. Combining data from the four nocodazole- and five colchicine-treated preparations, we found AVT also increased G significantly (P < 0.02) from 0.48 ± 0.07 to 0.70 ± 0.05 mS/cm2.
To determine whether Cl
movement might be responsible for the initial, AVT-stimulated,
transient that was abolished by microtubule disruption, we conducted
experiments in which most of the
Cl in the medium was
replaced by gluconate
(low-Cl Ringer). As shown
in Fig. 5, in both control and
low-Cl Ringer, AVT elicited
a two- to threefold increase in
Isc over a 1-h
period; however, the response to AVT of control cells was more rapid and showed an early transient peak that was not observed in
low-Cl Ringer in any of the
experiments in this series (Fig. 6). The basal Isc in
low-Cl Ringer (2.1 ± 0.9 µA/cm2) was significantly
lower (P < 0.01) than the
Isc in the
controls (6.8 ± 1.4 µA/cm2). In control cells AVT
significantly increased
Isc to 19.7 ± 3.0 µA/cm2
(P < 0.01), whereas
Isc in the
low-Cl Ringer increased to
5.4 ± 1.5 µA/cm2
(P < 0.05). After addition of AVT,
the conductance of the monolayers also increased significantly
(P < 0.02), from 0.52 ± 0.05 to
0.75 ± 0.04 and 0.18 ± 0.02 to 0.29 ± 0.04 mS/cm2, respectively, in the
control and low-Cl Ringer
preparations. In summary, the magnitude and time course of
AVT-stimulated
Isc recorded from
monolayers bathed in the
low-Cl Ringer was similar
to that recorded from monolayers treated with colchicine or nocodazole.
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The effect of nocodazole on the AVT-stimulated
Cl secretion was then
examined directly by experiments in which amiloride was used to block
the Na+ current and DIDS was added
to block Cl channels to
isolate the CFTR-mediated
Cl pathway. As shown by the
representative experiment in Fig. 7, nocodazole inhibited the increase in DIDS-insensitive and
amiloride-insensitive Isc produced by
AVT. As expected, basal
Isc was markedly
reduced by the presence of both amiloride and DIDS. In four experiments such as shown in Fig. 7, mean basal
Isc was 1.3 ± 0.6 µA/cm2 for control
(amiloride + DIDS treatment) and 1.2 µA/cm2 for preparations that
were also treated with nocodazole. However, in control monolayers, AVT
significantly increased
Isc by 0.33 ± 0.09 µA/cm2 (mean paired
difference, P < 0.05) in control but
not in nocodazole-treated monolayers (mean paired difference, 0.02 ± 0.04 µA/cm2).
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Because the preceding results suggested that the initial
Isc response to
AVT was due to net Cl
secretion, we examined the effect of glibenclamide, which has been
shown to inhibit CFTR (27), and
Cl replacement on the
amiloride-insensitive
Isc. A6
monolayers were treated with amiloride to block the
Na+ component of
Isc. As shown in
the experiment in Fig. 8, glibenclamide and
the low-Cl medium abolished
the stimulatory effect of AVT on the amiloride-insensitive Isc. In six
experiments such as that shown in Fig.
8B, AVT increased Isc significantly
from 0.5 ± 0.5 to +1.5 ± 0.7 µA/cm2
(P < 0.001 by paired
t-test). In the presence of
glibenclamide, the mean paired difference in
Isc (with vs.
without AVT) was 0.7 ± 0.2 (not significant), and in the
low-Cl medium it was
0.3 ± 0.1 µA/cm2 (not
significant). On the basis of these observations and the discussion
below, we hypothesized that CFTR mediated, at least in part, the
AVT-stimulated Cl secretion
present in A6 cells and that this
Cl secretion was inhibited
by microtubule disruption or glibenclamide.
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RT-PCR was performed on RNA obtained from A6 cells to probe for the presence of CFTR message. The PCR primers, which had been designed to amplify a portion of the CFTR R-domain based on the known Xenopus sequence, yielded a product of the size predicted by the Xenopus CFTR, 792 bp (Fig 9). This product was cloned and sequenced. The nucleotide and deduced amino acid sequences of this product were then compared with sequence information available for CFTR cloned from Xenopus (20) and A6 cells (4). Interestingly, the nucleotide sequence of the PCR product was more like the CFTR clone from Xenopus (98.6% nucleotide identity) than the clone from A6 cells (93.8% identity). The same was also true for the deduced amino acid sequence (97.7% identity with the Xenopus sequence and 92.7% identity with the A6 sequence).
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Immunocytochemistry on permeabilized A6 cells using an antibody prepared against the biochemically purified Na+ channel complex from bovine medulla (30) presented a pattern of apical, but not deep, intracellular labeling either before or after AVT treatment (Fig. 10). En face views (Fig. 11) localizing Na+ channels to the subapical and apical membrane region before and after AVT show no noticeable difference in staining. Fluorescence intensity of the apical membrane region was quantified in control and AVT-stimulated monolayers that had been processed for Na+ channel immunofluorescence. There was no significant difference in fluorescence intensity after AVT stimulation (0.159 ± 0.035 in control vs. 0.178 ± 0.042 after AVT, arbitrary fluorescence units, n = 10). There was, however, clear intracellular labeling with anti-Na+ channel antibody in cells that were subconfluent and not yet differentiated (Fig. 12), as well in confluent cells from a much later passage that never fully polarized (Fig. 13). Immunocytochemistry on permeabilized A6 cells utilizing an anti-CFTR antibody demonstrated both apical and intracellular localization of an immunoreactive protein. Furthermore, it appeared that there was a redistribution of the intracellular CFTR labeling to a more apical plane upon treatment of the monolayers with AVT (Fig. 14).
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Finally, the effect of nocodazole on the immunocytochemical localization of Na+ channels and CFTR in A6 cell monolayers was examined. Immunofluorescent localization with an anti-Na+ channel antibody revealed some intracellular labeling and a reduction of apical staining in nocodazole-treated cells compared with the untreated control (Fig. 15, left). In monolayers treated with nocodazole and stimulated with AVT, there was no noticeable effect on either the apical labeling or intracellular distribution of Na+ channel immunofluorescence compared with the nocodazole-treated cells (Fig. 15). Immunolocalization of CFTR in A6 cells treated with nocodazole shows diminished apical and disorganized intracellular labeling compared with the non-nocodazole-treated control (Fig. 15, right). Additionally, CFTR immunolocalization revealed that nocodazole prevented the AVT-stimulated redistribution of CFTR seen in Fig. 12 in the absence of nocodazole.
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DISCUSSION |
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Top
Abstract
Introduction
Methods
Results
Discussion
References
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We interpret the results presented above to reveal several important
features of the increase in
Isc produced in
A6 cells by AVT. First, AVT produces a biphasic change in
Isc that is the resultant of an early transient increase in
Cl secretion mediated by
CFTR and a slower but sustained increase in the amiloride-sensitive
sodium conductance (ASSC). Second, the early increase in CFTR-mediated
Cl transport depends on
intact microtubules and is associated with a movement of CFTR from the
deep cytoplasm to the apical region of the cells. In contrast, the late
increase in Isc
produced by ASSC occurs even when the microtubules have been disrupted
and is not associated with redistribution of ASSC from the cytoplasm to
the cell membrane surface. In the following discussion, we address
first the electrophysiological evidence in favor of these conclusions,
and second, we discuss the immunohistochemical evidence.
Electrophysiological evidence supporting an
AVT-stimulated trafficking of
Cl channels but not
Na+
channels.
We have presented the time courses for 10 control experiments (Figs. 3,
4, and 6), each of which demonstrates the biphasic nature of the
response to AVT. In the first 15-25 min after hormone addition,
there is a rise in
Isc that reaches
a peak and then declines to a stable or slowly decreasing level that we
refer to as the secondary phase. In the secondary phase,
Isc is completely inhibited by amiloride, indicating that all of the current in this
phase is mediated by the ASSC. In contrast, in nine preparations in
which the microtubules had been disrupted by treatment with nocodazole
(Fig. 3) or with colchicine (Fig. 4), the initial transient increase in
Isc was absent,
and there was only a slower and monotonic increase in
Isc that was
completely inhibited by amiloride.
secretion
mediated in part by CFTR. First, when all but 5 mM of the
Cl in the medium was
replaced with gluconate, the initial
Isc peak was
abolished (Figs. 5 and 6). This result is in agreement with the study
of Yanase and Handler (34), who previously described a secretory
Cl conductance in A6 cells
that had approximately the same time course. Second, when the ASSC was
blocked with amiloride, and DIDS was present to block other
Cl channels, pretreatment
with nocodazole prevented the increase in
Isc observed upon
AVT addition (Fig. 7). Third, when amiloride was again used to remove
the ASSC-mediated component of the response to AVT, there was still a
significant increase in
Isc upon AVT addition (Fig. 8B), but this was
prevented by glibenclamide (Fig. 8A)
or the low-Cl Ringer (Fig.
8C). Although glibenclamide is an
inhibitor of CFTR, it also inhibits ATP-sensitive
K+ channels (27). Such
K+ channels are present in A6
cells and may significantly contribute to the membrane potential (1);
therefore, it might be argued that the inhibition of
Cl secretion by
glibenclamide is not a conclusive proof of the involvement of CFTR.
To obtain more direct evidence for the presence of CFTR in these A6
cells, we used RT-PCR to demonstrate the presence of the CFTR message
(Fig. 9), but interestingly, the sequence of the PCR product obtained
was closer to the Xenopus sequence
than the A6 CFTR sequence recently published by Price et al. (24).
However, given the lack of specific selection pressure for CFTR in
cultured cells, it is not surprising that random mutations could have
resulted in variable divergence of the CFTR sequence from the parent
Xenopus sequence. In addition to the
evidence for CFTR mRNA in our A6 cells, immunofluorescence with an
anti-CFTR antibody detected an immunoreactive protein that
redistributed to the apical membrane after stimulation with AVT (Figs.
14 and 15). The latter results are discussed in more detail in the
following section on the immunofluorescence results.
Our conclusions regarding the transport events responsible for the
transient response of
Isc to AVT in A6
cells at first appear to be in direct contradiction to those of Verrey
et al. (33), who concluded that microtubule disruption in A6 cells
resulted in a 60% inhibition of ADH-stimulated ASSC but no inhibition
of ADH-stimulated Cl
conductance. However, we argue that we and Verrey et al. (33) are
drawing our conclusions from different time points in the response to
AVT. Our argument depends on a careful comparison of our results with
those of Verrey et al. (33) and others. For this purpose, we developed
Fig. 16, which is a composite of the time
courses of the
Isc response to
AVT (or AVP) in the present studies and those previously published by
Verrey et al. (33), Chalfant et al. (8), and Bindels et al. (2). To
facilitate the comparison, the time axes in the original studies were
expanded or contracted linearly to equalize the time scales, and
all four time courses were aligned vertically on the point of AVT/AVP
addition.
|
secretion. In our
present experiments, we have observed this very early transient only 7 times in 38 experiments (e.g., see experiments
2 and 3 in Fig. 6),
and it was not reported in the studies of Bindels et al. (2) or Yanase
and Handler (34). The reason for the variability is not clear, but it
is interesting to speculate that the
Cl secretion occasionally
seen within 3-5 min of AVT addition could represent activation of
CFTR residing in the apical membrane, whereas the
Cl secretion that occurs
subsequently (see below) involves trafficking of CFTR into the
membrane.
There is a local maximum (point B,
Fig. 16) that appears within 15-25 min after AVT/AVP addition in
all of the studies shown, as well as that of Yanase and Handler (34).
We have presented evidence that this local maximum is due, at least in
part, to Cl secretion, in
agreement with Yanase and Handler (34), and that this
Cl conductance is not
observed after microtubule disruption. Chalfant et al. (8) also
demonstrated that AVP-stimulated "oscillations" in
Isc, one of which
would correspond to point B in Fig.
16, disappeared in
Cl-free Ringer. Although
not emphasized by Chalfant et al. (8), this observation provides
additional evidence that point B in Fig. 16 is partially composed of a secretory
Cl current. In contrast,
Verrey et al. (33) described the effect of microtubule disruption on a
Cl conductance that
occurred immediately after AVT addition, whereas we are describing here
the effects of microtubule disruption on the
Cl conductance that occurs
15-25 min after AVT addition. Because the experiments of Verrey et
al. (33) ended within this later period, they did not reveal the second
and more prominent transient. Thus the discrepancy between our
conclusions and those of Verrey et al. (33) with regard to the effects
of microtubule disruption on
Cl conductance occur
because we are describing different points in the time course of the
response of Isc
to AVT. In the next section, we consider these differences in
interpretation in the light of changes in the cellular pattern of CFTR
and ASSC distribution with AVT.
Immunocytochemical support for an AVT-stimulated
trafficking of Cl channels but
not Na+
channels.
The immunocytochemistry results are most interesting when one compares
the localization of the Na+
channel and CFTR. First, in agreement with previous results (30), we
saw no intracellular binding of the
Na+ channel antibody except in
subconfluent cultures. In contrast, the fluorescent labeling of CFTR
was primarily intracellular, even in confluent, fully polarized
monolayers. Second, in the CFTR studies, there was a redistribution of
the intracellular staining to the apical membrane region after AVT
treatment. These results correlate nicely with observations of CFTR
"shuttling" in other epithelia (3, 12) and also with the
AVT-stimulated secretory Cl
conductance observed in our
Isc measurements.
It can also be seen in Fig. 15 that nocodazole inhibits the
AVT-stimulated translocation of CFTR.
secretion in A6 cells.
However, on the basis of the present data, it is difficult to determine
whether nocodazole inhibits
Cl secretion because of an
acute inhibition of membrane trafficking that normally regulates the
channel or whether the effect is more indirect, inhibiting perhaps the
total pool of Cl channels
within the biosynthetic pathway. Figure 15 shows that nocodazole
produced diminished apical labeling of CFTR and a general disorganization of the intracellular labeling. If one compares Fig. 15
with Fig. 14, it is clear that nocodazole inhibits AVT-dependent CFTR
translocation to the plasma membrane. This finding is in agreement with
Fuller et al. (13), who demonstrated that intact microtubules were
necessary for forskolin-stimulated
Cl secretion in T84 cells,
and with the observation of Toussan et al. (31) that nocodazole
inhibited forskolin-induced CFTR translocation to the plasma membrane
in T84 cells. Given the larger background of disorganized intracellular
CFTR labeling in the nocodazole-treated cells in Fig. 15, it is
difficult to conclude that translocation of CFTR with AVT was
completely eliminated. Nevertheless, if one compares Figs. 10 and 14,
it is clear that there is a distinctly different pattern of labeling
between Na+ channels and CFTR in
control cells, that only the pattern of CFTR labeling is clearly
altered by AVT, and that the latter alteration is prevented by
nocodazole (Fig. 15).
The absence of any effect of microtubule disruption on
Na+ transport in our studies is
consistent with the view that AVT may activate
Na+ channels already resident in
the plasma membrane. In support of this view, Oh et al. (21) have
demonstrated in vivo phosphorylation of
Na+ channel complexes in A6 cells
after stimulation with AVT. Furthermore, in vitro activation by PKA and
ATP of biochemically purified Na+
channel complexes from either A6 cells or bovine renal medulla alters
the kinetics of purified Na+
channels in a manner consistent with macroscopic
Na+ reabsorption (15, 21).
Membrane trafficking in response to AVT may, however, be dependent on
the developmental stage of the epithelium. Our immunofluorescence results show intracellular structures labeled with
anti-Na+ channel antibody in
subconfluent A6 cultures (Fig. 12) and in confluent cells before full
polarization has occurred (Fig. 13), but this perinuclear pattern of
labeling disappears as the monolayers "mature" (i.e., become
fully polarized). It is possible that in the early stages of
development AVT hastens the delivery of
Na+ channels to the apical surface
but that this ability to act as a "maturation factor" is lost as
polarity is fully achieved.
Our conclusions must also take into account the fact that the
immunofluorescence resolution is insufficient rule out shuttling of
Na+ channels between the cell
membrane and the cytoplasmic region immediately underlying it. In other
words, AVT might result in docking and insertion of
Na+ channels that are already
in close proximity to the apical membrane but that cannot be
distinguished from channels resident in the membrane. Such a mechanism
would be consistent with the observations of Kleyman et al. (16) and
Marunaka and Eaton (20), which indicate that AVT increases the absolute
number of Na+ channels in the
apical membrane of A6 cells. Furthermore, neither the bilayer results
cited above nor the present results clearly exclude membrane
trafficking as an regulatory mechanism that acts in addition to
activation of resident Na+
channels. On the other hand, if
Na+ channel shuttling occurs, then
the mechanism must be quite different than what is observed with CFTR,
which is translocated from the perinuclear cytoplasm to the apical
membrane. If Na+ channels
immediately underlying the apical membrane dock and fuse with that
membrane in response to AVT, then electron microscopy would be required
to resolve the process, but suitable antibodies for immunolocalization
at this level are not presently available.
| |
ACKNOWLEDGEMENTS |
|---|
We thank Christie Brown, M. L. Watkins, and B. C. Corbitt for expert technical assistance. In addition, we thank the many others who have contributed so much to this project: Drs. T. Howard (Univ. of Alabama) for rhodamine-phalloidin, S. Binder (Northwestern Univ.) for anti-tubulin antibody (Tu27-B), and C. Fuller, T. Wilborn, M. DuVall, C. Venglarik, M. Awayda, R. Rick, R. LeBoeuf, E. Schlatter, and K. Kirk for helpful discussions.
| |
FOOTNOTES |
|---|
This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grants DK-37206 and DK-25519.
Address for reprint requests: R. G. Morris, Dept. of Physiology and Biophysics, 958 MCLM Bldg., 1918 Univ. Blvd., Birmingham, AL 35294-0005.
Received 15 July 1996; accepted in final form 22 October 1997.
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REFERENCES |
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Top
Abstract
Introduction
Methods
Results
Discussion
References
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