cAMP-sensitive endocytic trafficking in A6 epithelia

Michael B. Butterworth1, Sandy I. Helman2, and Willem J. Els1

1 Department of Anatomy and Cell Biology, Faculty of Health Sciences, University of Cape Town, Observatory 7925, Cape Town, South Africa; and 2 Department of Molecular and Integrative Physiology, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Blocker-induced noise analysis and laser scanning confocal microscopy were used to test the idea that cAMP-mediated vesicle exocytosis/endocytosis may be a mechanism for regulation of functional epithelial Na+ channels (ENaCs) at apical membranes of A6 epithelia. After forskolin stimulation of Na+ transport and labeling apical membranes with the fluorescent dye N-(3-triethylammoniumpropyl)4-(6-4 diethylaminophenyl) hexatrienyl pyridinium dibromide (FM 4-64), ENaC densities (NT) decreased exponentially (time constant ~20 min) from mean values of 320 to 98 channels/cell within 55 min during washout of forskolin. Two populations of apical membrane-labeled vesicles appeared in the cytosol within 55 min, reaching mean values near 18 vesicles/cell, compared with five vesicles per cell in control, unstimulated tissues. The majority of cAMP-dependent endocytosed vesicles remained within a few micrometers of the apical membranes for the duration of the experiments. A minority of vesicles migrated to >5 µm below the apical membrane. Because steady states require identical rates of endocytosis and exocytosis, and because forskolin increased endocytic rates by fivefold or more, cAMP/protein kinase A acts kinetically not only to increase rates of cycling of vesicles at the apical membranes, but also principally to increase exocytic rates. These observations are consistent with and support, but do not prove, that vesicle trafficking is a mechanism for cAMP-mediated regulation of apical membrane channel densities in A6 epithelia.

epithelial sodium channel; noise analysis; confocal microscopy; channel trafficking; vesicles; forskolin; kidney; cortical collecting ducts; endocytosis


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

THE TRANSPORT OF SODIUM through amiloride-sensitive epithelial Na+ channels (ENaCs) is a major determinant of electrolyte and fluid homeostasis in mammals and other vertebrates. When expressed in native epithelia, absorption of Na+ at the apical membranes of the cells is determined not only by the electrochemical potential differences driving Na+ into the cells but also by the density of ENaCs that are in their functionally open state. Consequently, dysfunction of one or more of the Na+ channel subunits in apical membranes of several ion-transporting epithelia leads to pathological conditions characterized by impaired salt and water balance, such as hypertension (21). Consistent with its central role in Na+ absorption, it would not be surprising to find that ENaC activity is controlled by multiple regulatory factors. Generally, control of transport through ENaCs may be achieved by two independent and not mutually exclusive mechanisms, namely, by altering the gating kinetics or open probability of the channels and/or by altering the number of active or functional channels in the apical plasma membranes of the epithelial cells. Among kinases, protein kinase A (PKA) is known to play a major role in regulation of Na+ transport in a variety of tight epithelia and is thought to stimulate transport by increasing the total number or density of functional channels at the apical surfaces of the cells (7, 11, 17, 18, 25, 27). However, the various steps whereby ENaC densities are regulated by PKA are unknown, and elucidation of the steps and mechanisms remains a long-standing problem (10).

The density of several plasma membrane transporters is determined by mechanisms that control membrane trafficking of the transporters and/or their components to apical membranes [see review by Bradbury and Bridges (3)]. It is well appreciated that PKA modulates the water permeability of several epithelia through mechanisms that regulate the intracellular trafficking of water channels (4, 23). Evidence from earlier experiments supported the idea that arginine vasopressin stimulates Na+ transport by promoting the recruitment of Na+ channels to epithelial apical membranes (13). This thesis has received further but indirect support from several other sources using different approaches (11, 12, 22, 27, 34). Hence, although direct unequivocal evidence is not available to support this thesis, it is generally believed that ENaC densities may be regulated by mechanisms that control the rate of recruitment of channels and/or its subunits to the apical membrane and their endocytic retrieval. Although the mechanisms involved in epithelial cells and Xenopus oocytes may not be the same, it has been demonstrated that specific ENaC subunits expressed in oocytes contain sequences that allow them to be retrieved from the plasma membrane by endocytosis. Thus modification of the subunits by cytoplasmic proteins or other factors could serve as a signal that would lead to their endocytic retrieval and provide a means of regulating ENaC density in oocytes (6, 14, 30) and Madin-Darby canine kidney tissues (31).

To determine whether in A6 epithelia there is a cAMP-dependent apical membrane turnover mechanism that could be utilized for trafficking of ENaCs and/or its subunits, we imaged in live cells the endocytic cAMP-dependent process with scanning laser confocal microscopy. The rate of appearance of endocytic vesicles derived specifically from apical plasma membranes was correlated with results from blocker-induced noise analysis performed in parallel to measure the time courses of change of the rates of Na+ transport, functional ENaC densities, single channel currents, and channel open probabilities. These experiments were done during washout periods in tissues that were pretreated with forskolin to stimulate transport and ENaC densities. Collectively, the results are consistent with and support the idea that cAMP/PKA increases exocytosis, thereby providing a possible mechanism for regulation of the density of apical membrane ENaCs and hence Na+ transport in A6 epithelia.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Tissues and materials. The A6 cell line is a well-established model of a high-resistance, Na+-transporting epithelium.1 All experiments were performed on cultured A6 cells between passages 90 and 115. Cell cultures were grown to confluence on Nunc culture flasks in a humidified incubator at 28°C and gassed with 2.5% CO2. The culture medium was a modified DMEM (Highveld Biologicals, Johannesburg, South Africa), supplemented with 1% penicillin, 0.6% streptomycin, and 10% fetal calf serum (FCS; Highveld Biologicals). The medium was also supplemented with 10 ml/l of a 3% stock solution of L-glutamine. From these, cells were subcultured onto 30-mm permeable Millicell HA tissue culture inserts (0.45-µm pore size; Millipore, Bedford, MA) in culture medium. After ~10-21 days, the monolayers were confluent, and cells characteristically exhibited transcellular resistances of ~2 kOmega · cm2 or higher and transepithelial voltages >20 mV. Cells from the same batch of tissues and with similar electrical properties were used for the noise and for the confocal experiments.

Electrical measurements. Confluent monolayers of A6 cells were carefully punched out of the insert cups and mounted in perfusion chambers designed for noise analysis. The tissues were constantly perfused with growth medium minus the FCS and antibiotics.

The pulse protocol method of blocker-induced noise analysis allowed us to determine single channel currents (iNa), channel open probabilities (Po), and functional channel densities (NT) during steady state and transient periods of Na+ transport. The theory and protocols of this method have previously been described in detail (19). The method depends on measurements of the current noise power density spectra and the fractional inhibitions of the macroscopic amiloride-sensitive short-circuit currents (Isc) at two concentrations of the weak Na+ channel blocker, 6-chloro-3,5-diaminopyrazine-2-carboxamide (CDPC; Aldrich Chemical, Milwaukee, WI) (B10 = 10 and B30 = 30 µM CDPC). The following is a brief outline of the method. Tissues were short circuited continuously with a low-noise voltage clamp, and the resultant Isc was allowed to stabilize for 1-2 h before 10 µM CDPC was introduced into the apical perfusing medium for the duration of the experiment. CDPC (10 µM) caused a rapid small inhibition of Isc that was allowed to recover by autoregulatory mechanisms to baseline values of Isc within 30 min. At intervals of 10 min during the control and subsequent experimental periods, the apical solution was switched ("pulsed") to an identical solution that contained 30 µM CDPC. At the end of each experiment, 100 µM amiloride was added to the apical solution to measure the amiloride-insensitive Isc.

Current noise power density spectra were determined at 10 and 30 µM CDPC at each time point. Corner frequencies (fc) and low-frequency plateaus (So) of the Lorentzians were determined by nonlinear curve fitting of the spectra. Blocker on and off rates (kob and kbo, respectively) were determined from the slopes and intercepts of rate concentration plots (2pi fc = kobB + kbo), and the blocker equilibrium coefficient, KB, was calculated from the quotient kbo/kob at each time point. Single channel currents were calculated in the usual way (i<UP><SUB>Na</SUB><SUP>B</SUP></UP> = [So(2pi fc)2]/[4I<UP><SUB>Na</SUB><SUP>B</SUP></UP>kobB]), and Po was determined from the fractional inhibition of the blocker-sensitive Isc according to the following relationship
P<SUB>o</SUB><IT>=</IT><FENCE><FR><NU><IT>1−</IT>(<IT>I</IT><SUP><IT>30</IT></SUP><SUB>Na</SUB><IT>/I</IT><SUP><IT>10</IT></SUP><SUB>Na</SUB>)</NU><DE><IT>B<SUP>30</SUP></IT>(<IT>I</IT><SUP><IT>30</IT></SUP><SUB>Na</SUB><IT>/I</IT><SUP><IT>10</IT></SUP><SUB>Na</SUB>)<IT>−B<SUP>10</SUP></IT></DE></FR></FENCE><IT>K<SUB>B</SUB></IT> (1)
Open channel densities (N<UP><SUB>o</SUB><SUP>10</SUP></UP>) at 10 µM CDPC were calculated as I<UP><SUB>Na</SUB><SUP>10</SUP></UP>/i<UP><SUB>Na</SUB><SUP>10</SUP></UP>, whereas No in the absence of blocker was calculated as N<UP><SUB>o</SUB><SUP>10</SUP></UP>(1 PoB10/KB). The total pool of functional channels in closed and open states in the absence of blocker (NT) was calculated as No/Po.

Confocal fluorescence microscopy. To visualize the intracellular movement of apical membrane vesicles, we performed real-time confocal imaging of apical membranes in monolayers of A6 cells. To follow endocytic events, cell membranes were stained with the fluorescent marker N-(3-triethylammoniumpropyl)-4-(6-4 diethylaminophenyl) hexatrienyl pyridinium dibromide (FM 4-64; Molecular Probes, Leiden, Netherlands). This very specific lipophyllic styryl dye is widely used to visualize the movement of cell membranes in a number of experimental models (1). This dye is known to fluoresce weakly in aqueous solutions but strongly when bound to plasma membranes, cannot be readily washed out of the apical membranes of A6 epithelia (see RESULTS), is membrane impermeable, and does not migrate or exchange between intracellular compartments (1).

To visualize the endocytic retrieval of apical membranes during a period of forskolin washout, tissues on culture inserts were studied in their control state or were prestimulated with 2.5 µM forskolin added to the basolateral solution in DMEM for 35 min, as in the electrophysiological experiments, before membrane labeling. Apical membrane labeling was done in cold media to inhibit membrane trafficking and to inhibit the spread of the label down the basolateral plasma membranes. The tissues were chilled by briefly washing in cold (4°C) DMEM before being transferred to a cold medium that contained 16 µM of FM 4-64 prepared in DMEM (4 mM stock solution in DMSO) and kept on ice for 30 min. After labeling, the tissues were transferred to DMEM at room temperature, washed, removed from the inserts, and placed in DMEM in an inverted position on a glass coverslip (31-mm-diameter, 0.15-mm-thick; Bio Physical Technologies, Baltimore, MD) in a specially designed chamber for viewing on the confocal microscope. This usually took between 2 and 5 min. The chamber was not perfused but was sealed at the bottom with a silicone O-ring to prevent leakage or membrane movement while imaging. Control tissues were kept in DMEM equilibrated with room air without any forskolin for 35 min before being labeled exactly the same way as the experimental tissues.

Confocal images were obtained using a Zeiss LSM 410 inverted single laser confocal imaging system equipped with a Zeiss ×63 Planachromat oil objective [numerical aperture (NA) 1.4] and a ×100 Plan Nuofluar objective (NA 1.3) used for viewing. Fluorescence was excited by illuminating the cells with a krypton-argon laser using the 568-nm line that was close to the optimal absorption wavelength of 559 nm for FM 4-64. Optimum images of each optical section were obtained using scan times varying between 8 and 64 s. Individual lines were averaged (2-4 times). Preliminary experiments demonstrated that under these conditions, tissue autofluorescence was undetectable, photobleaching was minimal, and we could effectively image cells for extended periods (typically up to 1 h).

Optical x-y sectioning was done at z-increments of 1 µm, and the Zeiss LSM 410 software permitted color coding of the depth of the vesicles below the apical surfaces of the cells (see Fig. 6). The depth of the vesicles below the apical surfaces was also viewed in the x-z plane of the cells.

Statistical analysis. All data are expressed as means ± SE. Statistical analyses were performed in the usual way using paired t-tests when appropriate. P < 0.05 was considered significant.


    RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

A representative strip-chart recording is shown in Fig. 1 to illustrate typical responses of the Isc to addition of and subsequent washout of 2.5 µM forskolin from the basolateral solution. After a control period during which the tissues were pulsed with CDPC at intervals of 10 min, forskolin caused, after a short delay of a few minutes, maximal increases of Isc within ~30-40 min from time 0 control values that averaged 4.83 ± 0.60 µA/cm2. A summary of the blocker-sensitive INa at the various time points of noise analysis shown in Fig. 2A indicated that the mean increase of INa, expressed as experimental/control values, averaged 1.72 ± 0.12 at 35 min. The effect of forskolin on the INa was completely reversible. After washout of forskolin from the basolateral solution, the INa decreased from 7.96 ± 0.74 µA/cm2 at 35 min toward control values within ~60 min (Figs. 1 and 2A). At 95 min (55 min after forskolin was removed from the solution), INa averaged 5.46 ± 0.74 µA/cm2, a value only slightly higher than the original mean baseline control value. Noise analysis was carried out during both washin and washout of forskolin at the time points indicated in these figures. The experiments were terminated after addition of 100 µM amiloride to the apical solution that yielded the amiloride-insensitive current that averaged 0.28 ± 0.24 µA/cm2.


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Fig. 1.   A typical strip-chart recording of the time-dependent changes of the short-circuit current (Isc) caused after addition of 2.5 µM forskolin to the basolateral solution and during its subsequent washout. 6-Chloro-3,5-diaminopyrazine-2-carboxamide (CDPC; 10 µM) was present in the apical solution except for the pulse intervals (normally 10 min), when the CDPC concentration was increased to 30 µM. Isc was 1.97 µA/cm2 at the end of the control period and was increased by forskolin to 4.71 µA/cm2 within 35 min. Eighty minutes after forskolin washout, the Isc returned to near the time 0 control value, at which time 100 µM amiloride was used to inhibit blocker-sensitive Na+ transport. The amiloride-insensitive Na+ current was 0.20 µA/cm2.



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Fig. 2.   Summary of the time-dependent changes of the blocker-sensitive Na+ current (INa), single channel currents (iNa), channel open probabilities (Po), and functional epithelial Na+ channel densities (NT) caused by 2.5 µM forskolin and its subsequent washout from the basolateral solution. Data points were normalized to the last control point (-5 min) and expressed as experimental/control values (means ± SE; n = 11).

Forskolin-sensitive changes in functional ENaC densities. Blocker-induced noise analysis revealed that forskolin stimulated Na+ entry into the cells, and the subsequent decrease of transport during washout of forskolin was due to reversible time-dependent changes of the functional channels within the apical membranes of the cells. From mean control values of 84.4 ± 12.4 channels/100 µm2, NT was increased by forskolin to 285 ± 43.6 channels/100 µm2 within 35 min, or, as indicated in Fig. 2D, 3.50 ± 0.37-fold above control values. Within 55 min after washout of forskolin, NT decreased to near control values or to 98.3 ± 23.0 channels/100 µm2, with a quasi-exponential decline of NT with a time constant of ~20.5 min (see Fig. 7).

The time-dependent changes of NT were accompanied by reversible changes of both single channel currents and channel open probabilities (Fig. 2). Control iNa and Po measured just before treatment of the tissues with forskolin averaged 0.28 ± 0.04 pA and 0.26 ± 0.02, respectively. Within 5 min of forskolin treatment of the tissues, both iNa and Po were significantly decreased below control values (Fig. 2, B and C). At 35 min, iNa and Po decreased to 81.5 ± 2.4% and 61.5 ± 2.6% of their respective control values. After forskolin washout, iNa and Po returned to control (Po) or to slightly above control values (iNa). Accordingly, despite relatively small changes of both iNa and Po compared with those of NT, stimulation of Na+ transport by forskolin and the subsequent return of transport to control values after washout of forskolin were due to time-dependent changes of NT that were considerably larger in magnitude that those of iNa and Po.

Forskolin caused no significant change of the blocker interactions with the Na+ channels. The mean control values of the rate coefficients were kob = 8.56 ± 0.59 radians/(s · µM), kbo = 281.6 ± 20.0 radians/s, and KB averaged 34.1 ± 3.0 µM. These values remained essentially unchanged during stimulation and washout of forskolin as in previous studies of frog skin (11), indicating that channels recruited by forskolin possessed similar CDPC blocker kinetics as those present in the apical plasma membranes of A6 cells during the control period.

Internalization of apical membranes after withdrawal of forskolin. Previous studies with scanning electron microscopy demonstrated that the apical surfaces of A6 cells are not flat but dome shaped and that the cells are of unequal height (range of 8.0-17.6 µm) for tissues grown under the same conditions of the present experiments (9). Accordingly, with a nonuniform topography of adjacent cells of uneven heights, and nonuniform locations of tight junctions and lateral intercellular spaces, planar optical sectioning of groups of cells was defined to begin at the first level of appearance of fluorescence at the apical membrane surface of the tallest cell in the section (Fig. 3). Although it is not possible to rule out absolutely that FM 4-64 labeling of apical membranes had not progressed beyond the tight junctions, optical sectioning indicated clearly that this dye labeled the apical membranes of the cells and that the dye was irreversibly bound to the apical membranes for the duration of the experiments.


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Fig. 3.   A schematic two-dimensional cross section of epithelial cells grown on a permeable substrate illustrates nonuniform topography of adjacent cells of uneven heights and nonuniform locations of tight junctions (TJ) and lateral intercellular spaces (LIS). Sections were imaged at z-increments of 1 µm beginning at 0 µm, defined at the apical surface of the tallest cell. It should be noted that differences in cell heights need to be factored into assessments of the distance of vesicles below the apical surfaces of the cells in sections that contain many cells.

Figure 4 shows the internalization of apical membranes labeled with FM 4-64 in control and experimental cells according to the protocols described in MATERIALS AND METHODS. Images shown in Fig. 4 were acquired 40 min after forskolin washout and at a depth of 5 µm from the first section. It is clear that in control cells, there is very little internalization of vesicles originating from the apical membranes. The small quantity of vesicles in control cells probably represents the endogenous endocytosis occurring continuously in physiologically active cells. In contrast, in cells that have been previously treated with forskolin, there is a relative abundance of membrane vesicles beneath the apical membrane at 40 min after forskolin washout.


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Fig. 4.   Confocal micrographs show the internalization of vesicles derived from apical plasma membranes in control tissues (a) and in experimental tissues (b) 40 min after forskolin washout. The sections shown were imaged 5 µm below the apical surface of the tallest cell.

Kinetics of trafficking. We wished to determine whether the internalization of apical membrane corresponded to the time-dependent decreases of NT determined in the noise experiments during forskolin washout. To obtain an insight into the kinetics of membrane trafficking, three-dimensional imaging was performed on optical sections taken at 5 min and then at 10-min intervals throughout the period of disappearance of channels from the apical membranes. This allowed us to observe all the vesicles originating from the apical membranes within the cells. The gallery of images presented in Fig. 5 illustrates the internalization of apical membranes over a period of 55 min after removal of forskolin. A significant increase in the appearance of vesicles was already apparent in the images obtained at ~5 min after commencing forskolin washout. Thereafter, there was a progressive increase in vesicle appearance throughout the 50- to 60-min experimental periods, but where the rate of appearance of vesicles decreased with time. In two experiments carried out for longer times, large numbers of vesicles were apparent even after 120 min, but at these times, photobleaching became a limiting factor in data acquisition. In the control cells, there were far fewer vesicles at every time point (see Fig. 6) in the vicinity of the apical membranes of the cells.


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Fig. 5.   Confocal micrographs are shown illustrating the time course of appearance of endocytic vesicles derived from the apical plasma membranes at the time points indicated (5-55 min) after forskolin washout and are representative of 8 such experiments. The images are overlays of nine 1-µm optical sections starting 1 µm below the surface of the tallest cell. The mean planar area occupied by each cell is 104 µm2.



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Fig. 6.   Depth analysis of three-dimensional stacks overlayed in the x-y plane where fluorescent signals were assigned colors according to the z-scales shown in a, c, e, and g. Note difference in z-scales. In a and e, images are shown of a control tissue at 10 and 60 min; c and g are images at 10 and 60 min after forskolin washout. In b, d, f, and h, the same respective data are shown in the x-z plane where the dashed lines are drawn 5 µm below the apical surface of the tallest cell and where the density of vesicles in the overlayed sections at incremental depths of 1 µm is shown in white.

To determine the intracellular location of labeled vesicles, a depth analysis was performed as indicated in Fig. 6 (control and forskolin washout). The location of the vesicles was color coded depending on the relative z-position of the signal in the reconstructed three-dimensional stack. It was clear that shortly after forskolin washout (10 min), the majority of vesicles were located within 1-2 µm of the apical membranes (Fig. 6, c and g). Even after 60 min of forskolin withdrawal, the bulk of the vesicles remained within a few micrometers of the apical surface. However, a significant proportion of vesicles progressively distributed deeper into the cells during the forskolin washout period. Hence, after 55 min, some vesicles were visible at depths >5 µm beneath the first optical section and in an area near the nucleus, suggesting the existence of two populations of vesicles; those that remain in the vicinity of the apical membranes and those that migrate deeper into the cells. The latter population of vesicles appeared to migrate to deeper depths in both control tissues and those pretreated with forskolin, suggesting that endocytosis of these vesicles occurred in the absence of cAMP-dependent changes of Na+ transport. Thus vesicles that remain in the vicinity of the apical membranes may likely be those involved in cAMP-mediated trafficking of channels between the cytosol and the apical membranes of the cells.

To obtain a quantitative appreciation of the rates of appearance of vesicles in control and forskolin-pretreated tissues, the average number of vesicles per cell or vesicle density at each time point was plotted for individual experiments as indicated in Fig. 7B and summarized as indicated in Fig. 7C. In control tissues, the baseline vesicle density increased essentially linearly with time and averaged 4.8 ± 0.7 vesicles at 55 min with corresponding endocytic rates (vesicle density per minute) of 0.086 ± 0.010 vesicles per cell per minute. In the forskolin-pretreated tissues, the initial rates were far greater than in control tissues, as indicated in Fig. 7, B and C. With increasing time of forskolin washout, the endocytic rates appeared to decline approaching those of the baseline endocytic rates. Minimum endocytic rates of the forskolin-pretreated tissues estimated from the data at 15 min (assuming linear initial rates) averaged 0.45 ± 0.06 and from the data at 5 min averaged 0.69 ± 0.06. Recognizing that the actual endocytic rates would be somewhat higher if earlier time points could be acquired, we estimate as a first approximation that forskolin increased the endocytic rates by at least fivefold above the baseline rates of endocytosis. Because at the steady state, rates of endocytosis and exocytosis must be the same, exocytosis must be similarly increased rather markedly by forskolin.


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Fig. 7.   Time course of disappearance (A) of functional channel densities from the apical membranes of the cells and appearance (B) of endocytosed vesicle densities in a control tissue (gray squares) and a nonpaired forskolin pretreated tissue (solid squares) after forskolin washout. The solid line in A is the least-square regression fit of an exponential time-dependent decline of NT with a time constant of 20.5 min. The solid lines in B are the best fit regressions, which for the control baseline data, are linear. For the forskolin-pretreated tissues in B and C, the solid lines are best fit empirically and according to TableCurve (SPSS, Chicago, IL) to an equation of the form y = a + bxc. The dashed line in B indicates the approximate endocytic rate of the forskolin-pretreated tissue before washout of forskolin. See text for minimum estimates of endocytic rates of forskolin-pretreated tissues that were calculated with data points at 5 and 15 min. C: summary of vesicle densities at the various time points from tissues where the number of measurements indicated at each time point in parenthesis was >= 3 and in tissues where at least 2 measurements were made at various time points in the same tissue. In a separate group of 9 tissues pretreated with forskolin, vesicle densities at 45-55 min after forskolin washout averaged 18.3 ± 1.1 vesicles/cell. Note that the endocytic rate (vesicles per cell per minute) of the forskolin-pretreated tissues decreases with time approaching 60 min, the baseline endocytic rate of unstimulated tissues. Parameters of the forskolin-pretreated fitted line were a = 0.0379, b = 1.2809, and c= 0.5980. Endocytic rates determined as the first derivative of this line at 1, 5, and 55 min were 0.77, 0.40, and 0.15 vesicles per cell per minute, respectively. The baseline rate was 0.086 ± 0.010 vesicles per cell per minute. Accordingly, the endocytic rate had decreased from about 9-fold at 1 min to <2-fold above baseline endocytic rates within 55 min of forskolin washout.

It is imperative to recognize for both control and forskolin-treated tissues at steady states that the rates of endocytosis and exocytosis must be the same so that the exocytic rates can be determined from measurements of the endocytic rates. In this regard, the minimum increases of endocytic and exocytic rates estimated as indicated above clearly indicate at the forskolin steady state that cAMP/PKA causes a steady-state increase in the rates of gain and loss or cycling of vesicles at the apical membranes of the cells. This observation is consistent with the fact that enzymes can only increase the rates of reactions. With an elevated dynamic balance of endocytosis and exocytosis, it would be clear that cAMP/PKA does not act alone by inhibiting endocytosis. Thus of the many possible steps linking the pool(s) of cytosolic vesicles with the apical membranes of the cells, it remains to be determined which steps in the exocytic process are rate limiting and which steps are enhanced by cAMP/PKA, leading to increased rates of exocytosis.


    DISCUSSION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

cAMP is unquestionably a major short-term regulator of apical membrane Na+ transport in tight epithelia, including cell-cultured A6 epithelia. Forskolin stimulation of adenylate cyclase results under the conditions of the present experiments to near maximal and stable increases of Na+ entry into the cells within ~40 min due to increases of the density of functional ENaCs within the apical membranes of the cells. Stimulation of transport is completely reversible upon withdrawal of forskolin due to time-dependent decreases of functional channel densities. The fundamental and classic problem that addresses the underlying origin and disposition of cAMP-related changes of channel densities cannot be answered completely and unequivocally at this time. Functional channels within the apical membranes might originate by activation of nonfunctional channels that are already resident within the apical membranes. Functional channels might originate by assembly of channel subunits that reside within the apical membranes into functional channels. Functional channels might also originate by vesicle trafficking of functional channels to the apical membranes and/or subunits for later assembly within the apical membranes. Among such possibilities, there is presently no way known to us to perceive unequivocally, which, among these mechanisms, the cells use, in particular, for cAMP regulation of Na+ transport to achieve changes of the functional channel densities. Such questions will not be resolved until specific markers become available to distinguish between functional channels, nonfunctional channels, and channel subunits both within the apical membranes and the cytosol of the cells. There is no question that at some time during the life cycle of a functional channel, the channel itself or its subunits must be inserted into the apical membranes where the mature functional channel permits Na+ to enter the cells. Given the short term and reversible cAMP-dependent changes of NT, it would seem clear, regardless of mechanism, that cAMP must act through processes already in place with time constants in the range of minutes to elicit increases (forskolin) or decreases (forskolin washout) of functional channel densities within the apical membranes of the cells.

Our purpose in the present series of experiments was to test and to examine directly, for the first time, whether cAMP-mediated changes of transport could be correlated with changes of vesicle endocytosis at the apical membranes of intact epithelial cells. We chose specifically to label only apical membranes of the cells and to determine whether the time course of appearance in the cytosol of vesicles derived from apical membranes changed in parallel to the changes of functional channel densities. It was readily apparent from the time rate of appearance of vesicles that the endocytic rates during forskolin washout paralleled the time rate of decrease of NT, suggesting, but not proving, that the endocytosed vesicles contained the channels that disappeared from the apical membranes. This point was made in Fig. 7. If we assumed that the channels disappeared by way of vesicle endocytosis, then as a first approximation, each vesicle contained on average ~10 channels per vesicle (assuming that all vesicles contained ENaCs). Indeed, it must be appreciated quite generally that at physiological rates of Na+ transport, the physiologically relevant densities or absolute number of functional channels involved in Na+ transport in A6 and other tight epithelia (2) is quite low, which as a consequence, imposes methodological limitations in quantifying the actual number of functional channels. In this regard, however, it was equally of interest to find that the cellular densities of endocytosed vesicles were also remarkably quite low but consistent with trafficking of small numbers of channels. If, indeed, the disappearance of apical membrane channels occurs by endocytosis of vesicles, then quite remarkably it becomes apparent that vesicles may contain more than one channel and as many as 10 channels (or more), if this thesis is correct. Compatible with this thesis are observations by patch clamp of clustering of functional channels at the apical membranes of the cells (27). Thus clustering of functional channels may at least, in part, be due to trafficking of single vesicles that contain numerous channels.

With regard to mechanism of action of cAMP/PKA in stimulating Na+ transport, it is notable that we observed relatively large forskolin-related increases of endocytosis, which, at the steady state, mandate similar increases of exocytosis. Increases of NT can arise by either stimulation of exocytosis or inhibition of endocytosis of vesicles containing channels and/or their subunits. Although the site(s) of PKA action in vesicle trafficking is presently unknown in living epithelial tissues, activation of trafficking must occur by changes of either or both of the overall rate coefficients governing exocytosis/endocytosis. In this regard, forskolin activation of endocytosis implies and requires activation of exocytosis, so that in stimulated tissues, cycling of vesicles at the steady state is markedly elevated above baseline values. If cAMP/PKA acts only to inhibit the endocytic rate coefficient and hence the endocytic rate, thereby elevating NT and transport, we would expect no change in cycling of vesicles, because the exocytic rate coefficient and hence the exocytic rate would remain unchanged. Since we observed marked increases in cycling rate, the data are compatible with the thesis that cAMP/PKA must act predominantly, if not alone, to increase the exocytic rate coefficient.

It is also of interest to note here, as elsewhere, that mechanically preformed patches of apical membranes of A6 epithelia do not respond in any way to cAMP, even though unpatched apical membranes in the same tissues respond normally with increases of functional channel densities (27). Apparently, formation of membrane patches interferes with or precludes activation of channel densities through cAMP-dependent mechanisms. On one hand, it would be easy to explain this observation if formation of patches disrupts cAMP-dependent vesicle trafficking of channels to the apical membrane. On the other hand, we cannot yet dismiss unequivocally the possibility that patch formation may in addition disrupt mechanisms responsible for activation/assembly of dormant channels and/or subunits within the apical membranes to their functional states. Nevertheless, mechanical disruption of this type suggests that vesicle trafficking is involved as an important component of cAMP-related regulation of functional channel densities.

It is not possible to know with confocal imaging the exact size of each vesicle because the observed fluorescent image diameters would greatly exceed the actual sizes of the vesicles. In one group of cells, the measured fluorescent image diameters ranged between 0.18 and 0.84 µm with a mean ± SD of 0.51 ± 0.21 µm (n = 20). The actual mean diameter of the vesicles would be considerably <0.5 µm. If we assumed that vesicle diameters were near 0.1 µm, with surface area of 3.14 · 10-10 cm2, then the loss of membrane area due to endocytosis of 22 vesicles/cell would be ~0.66% of planar area (and less for actual membrane area) and correspondingly greater if vesicle diameters were 0.3 µm (5.9% of planar area). If vesicle diameters are, in fact, near or <0.1 µm, it would be either impossible with presently available methods or difficult, at best, to measure changes of membrane area either topographically or electrically through changes of membrane capacitance, or to find and identify such vesicles by electron microscopy. If vesicle diameters are >0.1 µm, it would be possible to detect changes of capacitance, especially if vesicle diameters were in the range of 0.3 µm or greater.2

The small number of vesicles that appear to be involved in transport may also impose limitations in the assessment of trafficking by use of fluid-phase markers, especially under circumstances where specific uptake by vesicles into the cells is small, relative to uncertainties imposed by nonspecific binding and/or uptake. Under favorable conditions, horseradish peroxidase (HRP) has been used as a fluid-phase marker. However, it has been recognized by others (20, 32) that HRP adsorbs onto plasma membranes and can be transported into cells by specific saturable mechanisms (35), thereby exacerbating determinations of specific vesicular uptake of this marker when background binding, sequestration, and/or other nonvesicular mechanisms of tissue uptake exceed or are in the range of vesicular uptakes.

Our findings, with regard to the action of forskolin on apical membrane endocytosis, are not the same as reported by Verrey et al. (33), who used HRP as a volume marker for endocytosis. Verrey et al. reported that neither aldosterone nor vasotocin or forskolin caused a change of apical membrane HRP uptake at 28°C (33). Background HRP activity was measured at 4°C, was assumed to be the same at 28°C, and was observed to be relatively high compared with total cell-associated activity that, according to these investigators, precluded measurements of short uptake periods. It is presently impossible to know what factor(s) preclude observation of forskolin-related endocytosis as measured by HRP uptake in nonaldosterone-treated tissues, especially when the increased endocytic rate is so readily observed by confocal microscopy as we reported in RESULTS.

We noted in RESULTS the existence of what appeared to be two populations of vesicles. One population was present in both control tissues and in tissues pretreated with forskolin. Characteristically, these vesicles were relatively few in number and could be observed to move into deeper sections (>5 µm) below the apical membrane surfaces of the cells at the later times of observation. The second population of vesicles observed during forskolin washout was more numerous, congregated just below the apical plasma membranes, and remained within 1-2 µm of the apical membrane for the 55-min periods of observation. Inspection of the confocal images at all time points indicated clearly that the subapical membrane vesicles were dissociated from the surface membranes (see footnote 2). It would be attractive to believe that these vesicles are likely to be involved in endocytic trafficking/cycling of channels because of their sustained presence in the vicinity of the apical membranes. Further speculation on possible (re)cycling of these vesicles and/or their final disposition is presently not warranted.

In summary, our finding of similar cAMP-mediated rates of disappearance of apical membrane functional channels and appearance of cytosolic vesicles originating from the apical membranes suggests that cAMP-mediated trafficking of channels and/or subunits of the channels in the A6 epithelium can occur through exocytosis/endocytosis. It will remain of particular interest to know to what extent channels can be packaged into vesicles for exocytosis and endocytosis and to what extent channel-packaging density in vesicles is a variable to be factored into understanding trafficking of channels and/or subunits to and from the apical membranes of the cells. Our findings, not only of relatively low physiological densities of membrane-bound functional channels but also of relatively low densities of vesicles involved in trafficking, underscore the need in experimental design and interpretation to consider the limits of detection of channels and vesicles involved functionally in regulation of ion transport at the apical membranes of tight epithelia.


    ACKNOWLEDGEMENTS

This work was supported by grants from the National Kidney Foundation of Southern Africa, the Medical Research Institute of South Africa, and the National Research Foundation (to W. J. Els) and by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-30824 (to S. I. Helman).


    FOOTNOTES

M. B. Butterworth is a doctoral student in the Department of Anatomy and Cell Biology, University of Cape Town Faculty of Health Sciences.

Address for reprint requests and other correspondence: S. I. Helman, Dept. of Molecular and Integrative Physiology, 524 Burrill Hall, 407 S. Goodwin Ave., Univ. of Illinois at Urbana-Champaign, Urbana, IL 61801(E-mail: s-helman{at}uiuc.edu).

1  In their hormone-unstimulated state, it is our understanding from our own experiments and those of others that A6 epithelia transport Na+ exclusively through their apical membranes regardless of the substrates, growth medium, serum, and other conditions of growth and regardless of their spontaneous rates of Na+ transport. Unlike classically studied toad urinary bladder and cortical collecting ducts where cAMP causes large increases of water permeability (15, 16, 29), the water permeability of A6 epithelia is insensitive to antidiuretic hormone (cAMP), indicating that in A6 epithelia, the apical membranes of the cells do not express water channels (8). Unlike toad urinary bladder and isolated preparations of frog skin (Rana pipiens) where cAMP does not change Cl- transport, cAMP increases Cl- transport in addition to Na+ transport in A6 epithelia (5, 36) by expression/activation of Cl- channels within the apical membranes of the cells (24, 26, 28). In this regard, it was of special interest during the course of the studies reported here to observe that the A6 epithelia grown under the conditions stated in MATERIALS AND METHODS did not respond to forskolin with an increase of Cl- transport. Noticeably absent in the Isc responses to forskolin were the immediate (<1 min) and unmistakable maximal increases of current that are characteristic of cAMP activation of Cl- current (Ref. 5 and S. I. Helman and T. Păunescu, personal communication) that precede the relatively slow secondary time course of activation of Na+ current. Consequently, although we do not know the factor(s) that govern expression of Cl- channels and the sensitivity or absence of sensitivity to cAMP, it was advantageous for the purpose of the present experiments to know insofar as presently possible that ion transport at the apical membranes was due principally, if not solely, to the transport of Na+.

2  Since completion of the work reported here, impedance analysis has been done to determine the time course of change of apical membrane capacitance in response to forskolin and its washout (T. Păunescu, W. Els, and S. I. Helman, personal communication) in A6 epithelia. The observation that capacitance changes timewise in parallel to the time-dependent changes of channel densities both during washin and washout of forskolin argues in support of the view that the labeled vesicles derived from the apical membranes observed during forskolin washout were completely dissociated from the apical membrane.

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Received 31 March 2000; accepted in final form 30 October 2000.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Betz, WJ, Mao F, and Smith CB. Imaging exocytosis and endocytosis. Curr Opin Neurobiol 6: 365-371, 1996[ISI][Medline].

2.   Blazer-Yost, BL, and Helman SI. The amiloride-sensitive epithelial Na+ channel: binding sites and channel densities. Am J Physiol Cell Physiol 272: C761-C769, 1997[Abstract/Free Full Text].

3.   Bradbury, NA, and Bridges RJ. Role of membrane trafficking in plasma membrane solute transport. Am J Physiol Cell Physiol 267: C1-C24, 1994[Abstract/Free Full Text].

4.   Brown, D, Katsura T, and Gustafson CE. Cellular mechanisms of aquaporin trafficking. Am J Physiol Renal Physiol 275: F328-F331, 1998[Abstract/Free Full Text].

5.   Chalfant, ML, Coupaye-Gerard B, and Kleyman TR. Distinct regulation of Na+ reabsorption and Cl- secretion by arginine vasopressin in the amphibian cell line A6. Am J Physiol Cell Physiol 264: C1480-C1488, 1993[Abstract/Free Full Text].

6.   Chalfant, ML, Denton JS, Langloh AL, Karlson KH, Loffing J, Benos DJ, and Stanton BA. The NH2 terminus of the epithelial sodium channel contains an endocytic motif. J Biol Chem 274: 32889-32896, 1999[Abstract/Free Full Text].

7.   Chou, KY, and Els WJ. Effects of disassembly of actin microfilaments on the AVP-induced regulation of sodium channel densities in frog skin epithelium. Biol Cell 89: 285-294, 1997[ISI][Medline].

8.   Crowe, WE, Ehrenfeld J, Brochiero E, and Wills NK. Apical membrane sodium and chloride entry during osmotic swelling of renal (A6) epithelial cells. J Membr Biol 144: 81-91, 1995[ISI][Medline].

9.   Els, WJ, and Butterworth MB. Cytochemical localization of adenylate cyclase in cultured renal epithelial (A6) cells. Microsc Res Tech 40: 455-462, 1998[ISI][Medline].

10.   Els, WJ, and Helman SI. Regulation of epithelial sodium channel densities by vasopressin signaling. Cell Signal 1: 533-539, 1989[ISI][Medline].

11.   Els, WJ, and Helman SI. Activation of epithelial Na channels by hormonal and autoregulatory mechanisms of action. J Gen Physiol 98: 1197-1220, 1991[Abstract].

12.   Fisher, RS, Grillo FG, and Sariban-Sohraby S. Brefeldin A inhibition of apical Na+ channels in epithelia. Am J Physiol Cell Physiol 270: C138-C147, 1996[Abstract/Free Full Text].

13.   Garty, H, and Edelman IS. Amiloride-sensitive trypsinization of apical sodium channels. Analysis of hormonal regulation of sodium transport in toad bladder. J Gen Physiol 81: 785-803, 1983[Abstract].

14.   Goulet, CC, Volk KA, Adams CM, Prince LS, Stokes JB, and Snyder PM. Inhibition of the epithelial Na+ channel by interaction of Nedd4 with the PY motif deleted in Liddle's syndrome. J Biol Chem 273: 30012-30017, 1998[Abstract/Free Full Text].

15.   Grantham, JJ, and Burg MB. Effect of vasopressin and cyclic AMP on permeability of isolated collecting tubules. Am J Physiol 211: 255-259, 1966[ISI][Medline].

16.   Hays, RM, and Leaf A. Studies on the movement of water through the isolated toad bladder and its modification by vasopressin. J Gen Physiol 45: 905-919, 1962[Abstract/Free Full Text].

17.   Helman, SI, Cox TC, and Van Driessche W. Hormonal control of apical membrane transport in epithelia. Studies with fluctuation analysis. J Gen Physiol 82: 201-220, 1983[Abstract].

18.   Helman, SI, Els WJ, Cox TC, and Van Driessche W. Hormonal control of the Na entry process at the apical membrane of frog skin. In: Membrane Biophysics: Structure and Function in Epithelia, edited by Dinno M, and Callahan A.. New York: Liss, 1981, p. 47-56.

19.   Helman, SI, Liu X, Baldwin K, Blazer-Yost BL, and Els WJ. Time-dependent stimulation by aldosterone of blocker-sensitive ENaCs in A6 epithelia. Am J Physiol Cell Physiol 274: C947-C957, 1998[Abstract/Free Full Text].

20.   Hew, WSR, Robertson AJ, Ross P, and Hopwood D. The study of the process of fluid-phase endocytosis in cervical squamous cells using fluorescent microspheres. Cytopathology 10: 375-382, 1999[ISI][Medline].

21.   Hummler, E, and Horisberger JD. Genetic disorders of membrane transport. V. The epithelial sodium channel and its implication in human diseases. Am J Physiol Gastrointest Liver Physiol 276: G567-G571, 1999[Abstract/Free Full Text].

22.   Kleyman, TR, Ernst SA, and Coupaye-Gerard B. Arginine vasopressin and forskolin regulate apical cell surface expression of epithelial Na+ channels in A6 cells. Am J Physiol Renal Fluid Electrolyte Physiol 266: F506-F511, 1994[Abstract/Free Full Text].

23.   Knepper, MA, and Inoue T. Regulation of aquaporin-2 water channel trafficking by vasopressin. Curr Opin Cell Biol 9: 560-564, 1997[ISI][Medline].

24.   Kokko, KE, Matsumoto PS, Zhang ZR, Ling BN, and Eaton DC. Prostaglandin E2 increases 7-pS Cl- channel density in the apical membrane of A6 distal nephron cells. Am J Physiol Cell Physiol 273: C548-C557, 1997[Abstract/Free Full Text].

25.   Li, JHY, Palmer LG, Edelman IS, and Lindemann B. The role of sodium-channel density in the natriferic response of the toad urinary bladder to an antidiuretic hormone. J Membr Biol 64: 77-89, 1982[ISI][Medline].

26.   Marunaka, Y, and Eaton DC. Chloride channels in the apical membrane of a distal nephron A6 cell line. Am J Physiol Cell Physiol 258: C352-C368, 1990[Abstract/Free Full Text].

27.   Marunaka, Y, and Eaton DC. Effects of vasopressin and cAMP on single amiloride-blockable Na channels. Am J Physiol Cell Physiol 260: C1071-C1084, 1991[Abstract/Free Full Text].

28.   Nakahari, T, and Marunaka Y. ADH-evoked [Cl-]i-dependent transient in whole cell current of distal nephron cell line A6. Am J Physiol Renal Fluid Electrolyte Physiol 268: F64-F72, 1995[Abstract/Free Full Text].

29.   Orloff, J, Handler JS, and Preston AS. The similarity of effects of vasopressin, adenosine-3'-5'-phosphate (cyclic AMP) and theophylline on the toad bladder. J Clin Invest 41: 702-709, 1962[ISI].

30.   Shimkets, RA, Lifton RP, and Canessa CM. The activity of the epithelial sodium channel is regulated by clathrin-mediated endocytosis. J Biol Chem 272: 25537-25541, 1997[Abstract/Free Full Text].

31.   Staub, O, Gautschi I, Ishikawa T, Breitschopf K, Ciechanover A, Schild L, and Rotin D. Regulation of stability and function of the epithelial Na+ channel (ENaC) by ubiquitination. EMBO J 16: 6325-6336, 1997[Abstract/Free Full Text].

32.   Stromhaug, PE, Berg TO, Gjoen T, and Seglen PO. Differences between fluid-phase endocytosis (pinocytosis) and receptor-mediated endocytosis in isolated rat hepatocytes. Eur J Cell Biol 73: 28-39, 1997[ISI][Medline].

33.   Verrey, F, Digicaylioglu M, and Bolliger U. Polarized membrane movements in A6 kidney cells are regulated by aldosterone and vasopressin/vasotocin. J Membr Biol 133: 213-226, 1993[ISI][Medline].

34.   Verrey, F, Groscurth P, and Bolliger U. Cytoskeletal disruption in A6 kidney cells: impact on endo/exocytosis and NaCl transport regulation by antidiuretic hormone. J Membr Biol 145: 193-204, 1995[ISI][Medline].

35.   Yamaguchi, Y, Dalle-Molle E, and Hardison WGM Hepatocyte horseradish peroxidase uptake is saturable and inhibited by mannose-terminal glycoproteins. Am J Physiol Gastrointest Liver Physiol 264: G880-G885, 1993[Abstract/Free Full Text].

36.   Yanase, M, and Handler JS. Adenosine 3',5'-cyclic monophosphate stimulates chloride secretion in A6 epithelia. Am J Physiol Cell Physiol 251: C810-C814, 1986[Abstract/Free Full Text].


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