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Am J Physiol Renal Physiol 274: F300-F314, 1998;
0363-6127/98 $5.00
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Vol. 274, Issue 2, F300-F314, February 1998

Microtubule disruption inhibits AVT-stimulated Clminus secretion but not Na+ reabsorption in A6 cells

Ryan G. Morris1, Albert Tousson2, Dale J. Benos1,2, and James A. Schafer1,3

Departments of 1 Physiology and Biophysics, 2 Cell Biology, and 3 Medicine, University of Alabama at Birmingham, Birmingham, Alabama 35294

    ABSTRACT
Top
Abstract
Introduction
Methods
Results
Discussion
References

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

    INTRODUCTION
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Abstract
Introduction
Methods
Results
Discussion
References

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.

    METHODS
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Abstract
Introduction
Methods
Results
Discussion
References

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 beta -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 beta -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, beta -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(beta -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).

Electrical measurements. VT of the cultures was monitored with an EVOM (World Precision Instruments, Sarasota, FL). Measurement of Isc (µA/cm2) and transepithelial conductance (G, mS/cm2) was done in a Ussing-type chamber, which has been described in detail by Rick et al. (25). The filter was mounted cell-side-up on a plastic ring in one half-chamber. The other half-chamber was then attached, clamping the filter in the middle of the Ussing-type chamber. The chambers were connected to a flow-through system for continuous recirculation of the solution on each side (1.0 ml/min) with a roller pump (Ismatec, Zurich, Switzerland). Chambers were kept in an incubator to maintain the temperature at 26°C, and solutions were continuously bubbled with 99% O2-1% CO2. Each half-chamber housed a potential-sensing Ag-AgCl electrode (IVM, Healdsburg, CA) and an Ag-AgCl current passing electrode. All electrodes were connected to a voltage clamp (model VCC600; Physiological Instruments, Houston, TX). The Isc was continuously recorded on a strip-chart recorder. G was determined at 90-s intervals by passing a 10-mV pulse of 1-s duration.

The standard solution used to perfuse the chambers contained (in mM) 100 NaCl, 1 CaCl2, 0.5 MgCl2, 2 K2HPO4, 10 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES), and 5 glucose. A "low-Cl- 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, beta -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.

    RESULTS
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Abstract
Introduction
Methods
Results
Discussion
References

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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  		Fig. 1.   Immunofluorescent staining of microtubules demonstrating effect of colchicine and nocodazole on microtubule structure. All are en face views. Bars = 10 µm.

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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  		Fig. 2.   Representative time course of arginine vasotocin (AVT)-stimulated short-circuit current (Isc) in presence or absence of nocodazole pretreatment. Downward deflections are the current change for a 10-mV change in transepithelial voltage (VT) and are proportional to the conductance. AVT was added at 0.1 µM, and amiloride was added at 10 µM (arrows).


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  		Fig. 3.   Results from three experiments in which the time course of Isc was measured in presence and absence of nocodazole as described for Fig. 2. For clarity, Isc deflections for conductance measurements have been omitted. In each of three experiments, one of the culture inserts was used as a control and another was pretreated with nocodazole. Number following the label for each curve is the Isc value in µA/cm2 immediately before AVT addition. Time of addition of 0.1 µM AVT is indicated by the vertical bar below each curve, and the time of addition of 10 µM amiloride is indicated by the bar above each curve. In the case of experiment 2, two inserts were treated with nocodazole (traces 2a and 2b); control and one nocodazole record (trace 2b) were shown in Fig. 2 with a more compressed time scale.


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  		Fig. 4.   Results from three experiments in which the time course of Isc was measured in presence and absence of colchicine following the same protocol as described for Fig. 3. Time of addition of 0.1 µM AVT is indicated by the first vertical bar, and the time of addition of 10 µM amiloride is indicated by the second vertical bar. In the case of experiments 2 and 3, two inserts were treated with colchicine (traces 2a, 2b, 3a, and 3c).

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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  		Fig. 5.   Effect of low-Cl- Ringer on the time course of Isc after addition of 0.1 µM AVT. Bracket labeled "2°" below the control curve denotes secondary phase of AVT response. Amiloride was added at 10 µM.


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  		Fig. 6.   Results from four experiments in which the time course of Isc was measured in normal medium or in low-Cl- Ringer. Time of addition of 0.1 µM AVT is indicated by the first vertical bar, and the time of addition of 10 µM amiloride is indicated by the second. Experiment 1 is the same experiment as shown with a compressed time scale in Fig. 5.

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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  		Fig. 7.   Effect of nocodazole on AVT-stimulated, DIDS-insensitive Cl- secretion. In both control and nocodazole-treated cells, Na+ current was inhibited by 10 µM amiloride. Amiloride addition was followed by 100 µM DIDS and then 0.1 µM AVT.

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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  		Fig. 8.   Effects of 300 µM glibenclamide or low-Cl- Ringer on AVT-stimulated Cl- secretion. Amiloride at 10 µM was present in all three inserts. A: glibenclamide also added at 300 µM. B: amiloride only. C: low-Cl- Ringer was used as the bathing solution.

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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  		Fig. 9.   Polymerase chain reaction amplification (30 cycles) of cystic fibrosis transmembrane conductance regulator (CFTR) from A6 cells utilizing primers designed to amplify the R-domain of Xenopus CFTR. MW, molecular weight standards.

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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  		Fig. 10.   Immunofluorescence localization of Na+ channels in A6 cell monolayers before (left) and after (middle) AVT stimulation (right, preimmune controls). All are side views, and arrowheads denote position of apical surface. A: corresponding phase-contrast micrographs. B: immunofluorescence localization of Na+ channels. C: Hoechst staining of nuclei. Bars = 20 µm.


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  		Fig. 11.   En face views demonstrating Na+ channel immunolocalization before (left) and after (right) vasotocin (AVT) stimulation. A: immunofluorescence of apical membrane. B: subapical immunofluorescence. C: nuclei. Bars = 20 µm.


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  		Fig. 12.   En face views demonstrating Na+ channel immunolocalization in subconfluent A6 cells before (left) and after (right) AVT stimulation. A: immunofluorescence localization of Na+ channels. B: Hoechst staining of nuclei. Bars = 20 µm.


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  		Fig. 13.   Na+ channel localization in partially polarized but confluent A6 monolayers. All are side views, and arrowheads denote apical surface. Bars = 20 µm.


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  		Fig. 14.   Immunocytochemical localization of CFTR in A6 cells before and after vasotocin (AVT) stimulation. Both control and experimental epithelia were taken from paired culture inserts in which VT was continuously measured. Approximately 15 min after addition of AVT, when VT had apparently reached a maximum and not yet declined, membranes were cut out of the plastic inserts, washed with ice-cold phosphate-buffered saline, and immediately fixed. All are side views. Top: immunofluorescence of CFTR. Bottom: Hoechst staining of nuclei. Bars = 20 µm.

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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  		Fig. 15.   Immunofluorescence of Na+ channels (left) and CFTR (right) demonstrating inhibitory effect of nocodazole on apical localization of channels before and after AVT stimulation. All are side views, and arrowheads denote apical surface. Bars = 20 µm.

    DISCUSSION
Top
Abstract
Introduction
Methods
Results
Discussion
References

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.

It is clear that Isc was diminished in the nocodazole- and colchicine-treated cells both before (basal Isc) and after AVT addition. Nevertheless, AVT produced a significant increase in Isc, despite the disruption of microtubules, and in these experiments AVT increased Isc by 2.0 (colchicine) to 3.9 (nocodazole) times the basal current, which is, if anything, greater than the doubling of the basal current produced in the control cells in the secondary phase. It seems most likely that the lower currents in cells with disrupted microtubules may be the result of an inhibition of the biosynthetic delivery of one or more transport proteins or their regulators, of a general inhibition of cellular metabolic processes, or both.

Three sets of electrophysiological experiments strongly suggest that the initial stimulation of Isc by AVT was due to Cl- 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.


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  		Fig. 16.   Composite of Isc tracings from Verrey et al. (33), Chalfant et al. (8), present data, and Bindels et al. (2) comparing the time course of the response of A6 cells to AVT or AVP. AVT or AVP doses were as follows: Verrey et al., 0.028 µM; Chalfant et al., 0.220 µM; present study, 0.1 µM; and Bindels et al., 0.1 µM. See text for explanation.

The time courses of Verrey et al. (33) and Chalfant et al. (8) are similar in that they demonstrate a transient increase in Isc immediately after AVT addition (point A in Fig. 16), which these investigators attribute to Cl- 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.

It is clear from the immunofluorescence studies of control monolayers that there is a distinctly different pattern of labeling between Na+ channels and CFTR, and only the pattern of CFTR labeling is clearly altered by AVT. Our results with the Na+ channel antibody (Fig. 10) indicate that Na+ channels are not acutely redistributed to the apical membrane from a cytoplasmic pool, at least not in highly differentiated cultures of A6 epithelia. Additionally, the immunolocalization of Na+ channels in the presence of nocodazole (Fig. 15) clearly shows a diminished apical labeling relative to control. This reduction in the density of apical Na+ channels may, in part, explain the diminished basal Isc found in nocodazole- or colchicine-treated cell. However, at least two other transporters needed for transepithelial Na+ transport (Na-K-ATPase and K+ channels) may also be inhibited by microtubule disruption. Despite the apparent decrease in apical Na+ channel density in the nocodazole-treated cells, there is still a proportional increase in AVT-stimulated Na+ transport.

These observations should be considered in the light of the conclusions reached by Verrey et al. (33) in comparison with our own. In this study and that of Verrey et al., there is agreement that microtubule disruption lowers basal current levels and also AVT-stimulated Na+ conductance. Furthermore, in neither study does microtubule disruption prevent activation of Na+ channels, so that AVT produces a proportional increase in ADH-stimulated current relative to the basal current values in the presence or absence of microtubule inhibitors. However, the study of Verrey et al. (33) did not take into account the possibility that microtubule disruption might diminish the absolute number of Na+ channels at the apical membrane via an inhibition of the biosynthetic delivery pathway. It is clear from the immunofluorescence data presented here that microtubule disruption does indeed diminish the apical labeling of Na+ channels relative to untreated controls. Thus the decrease in ADH-stimulated Na+ transport reported by Verrey et al. (33) is most likely due to the diminished number of transporters in the cell membrane rather than an acute inhibition of membrane trafficking. AVT would still activate the Na+ channels in the presence of microtubule disruption but only in proportion to the number of channels that are still resident in the apical membrane.

The same argument might apply to the effects of nocodazole on Cl- 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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Abstract
Introduction
Methods
Results
Discussion
References

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