Department of Neuroscience and Physiology, State University of New York Upstate Medical University, Syracuse, New York 13210
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Whole cell
patch-clamp techniques were used to investigate amiloride-sensitive
sodium conductance (GNa) in the everted initial collecting tubule of Ambystoma. Accessibility to both the
apical and basolateral membranes made this preparation ideal for
studying the regulation of sodium transport by insulin.
GNa accounted for 20% of total cell conductance
(GT) under control conditions. A resting
membrane potential of 75 ± 2 mV (n = 7)
together with the fact that GT is stable with
time suggested that the cells studied were viable. Measurements of
capacitance and use of a known uncoupling agent, heptanol, suggested
that cells were not electrically coupled. Thus the values of
GT and GNa represented individual principal cells. Exposure of the basolateral membrane to
insulin (1 mU/ml) for 10-60 min significantly (P < 0.05) increased the normalized GNa [1.2 ± 0.3 nS (n = 6) vs. 2.0 ± 0.4 nS
(n = 6)]. Cell-attached patch-clamp techniques were
used to further elucidate the mechanism by which insulin increases
amiloride-sensitive epithelial sodium channel (ENaC) activity. In the
presence of insulin there was no apparent change in either the number
of active levels/patch or the conductance of ENaC. The open
probability increased significantly (P < 0.01) from
0.21 ± 0.04 (n = 6) to 0.46 ± 0.07 (n = 6). Thus application of insulin enhanced sodium reabsorption by increasing the fraction of time the channel spent in
the open state.
sodium conductance; epithelial sodium channel
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
FINE CONTROL OF SODIUM REABSORPTION occurs in the distal nephron, where sodium homeostasis is achieved via regulation of the rate of its reabsorption. Sodium reabsorption is a two-step process, the entry of sodium on the apical side and the extrusion from the basolateral side (46). The Na+-K+-ATPase located in the basolateral membrane extrudes the intracellular sodium, maintaining the electrochemical gradient for the apical entry of sodium. On the apical side, sodium enters the cell through the amiloride-sensitive epithelial sodium channel (ENaC).
The highly selective 5-pS ENaC, comprised of three subunits (,
,
and
), is the predominant sodium channel found in the distal nephron
(8, 17, 18, 30-32, 42, 44). Mutations in ENaC can
lead to salt-retaining or salt-wasting disorders (18). These observations have led to the general view that ENaC is
responsible for most, if not all, of the fine tuning of sodium
reabsorption in the late distal nephron. Although other sodium-carrying
channels have been reported in cell lines derived from various
epithelial cells (for review, see Ref. 18) and in
collecting tubules of potassium-adapted amphibia (45),
they have not been observed in native rat or salamander collecting
tubules under control conditions, making evaluation of their
significance to sodium homeostasis difficult.
There is a considerable body of literature on the modulation of ENaC by hormones such as aldosterone and vasopressin. However, little is known about the regulation of sodium reabsorption by insulin, a hormone known to increase sodium reabsorption in the mammalian kidney (11, 12, 16) and amphibian model systems of the distal nephron (2, 5, 6, 9, 15, 27, 34). Increases in sodium reabsorption by insulin could be caused by several factors: 1) an increase in the open probability of the channel, 2) an increase in the conductance of ENaC, or 3) an increase in the number of active channels. The latter may represent the activation of quiescent ENaCs already present in the membrane or the synthesis and/or insertion of ENaCs into the membrane from a cytosolic pool.
The mechanism(s) by which insulin regulates sodium balance is controversial. Marunaka et al. (27) used a cell-attached patch-clamp technique on A6 cells to demonstrate that insulin increases the open probability of ENaC without any increase in active channel number. However, investigators using noise analysis techniques show no increase in ENaC open probability but do report an increase in the number of active channels in the apical membrane of A6 cells (2, 6, 15). It is unclear whether the different findings are due to the use of tissue-cultured epithelial cells, electrophysiological methods, or duration of insulin exposures.
The ability of insulin to increase sodium reabsorption occurs at
physiological concentrations that are independent of changes in
glucose, aldosterone, or vasopressin (10). Plasma insulin levels are known to be elevated in clinical conditions such as diabetes
mellitus type II or obesity. In the obese Zucker rat, a model for type
II diabetes, the expression of the -subunit of ENaC is upregulated
in rats at 2 and 4 mo of age (3). It is not known whether
the increase in this subunit correlates to an increase in sodium
reabsorption. It is possible that over time (weeks, months, or years),
the effects of the hyperinsulinemic condition on ENaC could increase
sodium reabsorption, expand extracellular volume, and lead to
hypertension (3, 10, 20).
In native renal tissue, the mechanism that insulin utilizes to increase sodium transport at the single-cell level is unknown. The distal tubule segment of the amphibian nephron is a model system of the mammalian distal nephron and shares many similarities in structure and function (22, 38-41). It is possible to evert the initial collecting tubule of the tiger salamander, Ambystoma tigrinum, and perform electrophysiological techniques on the apical membrane while separately controlling the contents of the saline bathing the apical and basolateral membranes (14, 42, 44, 45). The everted tubule is ideal for studying the effects of hormones on the cellular mechanisms of sodium reabsorption.
A whole cell patch-clamp technique was applied to the apical membrane of the everted initial collecting tubule to characterize some of the electrophysiological properties of principal cells. We found the resting membrane potential to be near the calculated equilibrium potential for potassium. The sodium conductance (GNa) was ~20% of total cell conductance (GT) and remained stable with time. These data indicate the viability of the principal cells of the everted initial collecting tubule.
We used both the whole cell and cell-attached patch-clamp techniques on everted initial collecting tubules treated with insulin in vitro. We demonstrate that insulin nearly doubled the whole cell sodium conductance. In cell-attached patch-clamp experiments, insulin caused a significant increase in the open probability of ENaC.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
General. Western variety, aquatic-phase Ambystoma tigrinum (Charles Sullivan, Nashville, TN) were housed in an aquarium at 55°F with circulating tap water and were fed crickets daily. Immediately before an experiment, a salamander was pithed and the kidney was rapidly removed. Several cross-sectional slices of kidney, 3-4 mm thick, were cut for dissection. Initial collecting tubules were dissected and trimmed to a length of 1-1.5 mm. The segment was then transferred to a modified renal tubule perfusion chamber that was mounted on the stage of an inverted microscope (Zeiss). To prevent the tubule from rolling during patch-clamp experiments, the bottom of the perfusion chamber was pretreated with Cell-Tak (Collaborative Research, Boston, MA).
Eversion and perfusion. The technique and experimental setup used to evert and perfuse kidney tubules were a modification of those developed by Burg et al. (7) and have been described in detail previously (14, 42, 43). Briefly, specialized glass pipettes were fabricated to turn the initial collecting tubule inside out, exposing the apical membrane (14, 42). To prevent the collecting tubules from sticking to the glass pipettes during the eversion process, these pipettes were soaked in a saline solution containing 1 g/100 ml albumin. A system of valves and reservoirs permitted the investigator to superfuse a variety of saline solutions over either the apical or the basolateral surface of the everted tubule. Via gravity feed, saline flowed continuously to exchange fluid in the inner perfusion pipette during the experiment, providing fluid to the basolateral surface. Both apical and basolateral flow rates were adjusted such that the exchange of solution was 95% complete within 1 min.
Patch-clamp methods.
Methods to fabricate patch-clamp pipettes and form seals for
cell-attached and whole cell patch-clamp studies were described previously (17, 21, 42-45). Pipettes were fabricated
from 100-µl Microcaps (Drummond Scientific, Broomall, PA) on a
Brown-Flaming P-80/PC puller (Sutter Instrument, San Rafael, CA)
immediately before use. The pipette tips were fire polished on a
Narishige microforge (Narishige, Tokyo, Japan). Pipette resistances
ranged between 2.5 and 3.3 M.
Whole cell recordings.
Whole cell voltage-clamp studies were performed by forming a gigaohm
seal on the apical membrane as described in Patch-clamp methods. Intrinsic capacitance of electrodes was
compensated on-line. The cell membrane was ruptured by applying suction
to the pipette until an abrupt decrease in resistance and increase in
capacitance was observed on the oscilloscope. The holding potential was
80 mV. Membrane potentials were stepped in 20-mV increments ranging from
100 mV to 0 mV with a pulse duration of 100 ms. The current at
each voltage was determined 30-40 ms after pulse onset.
GT was determined from the slope of the
current-voltage plot in the linear range of
100 to 0 mV. At the end
of the experiment, withdrawal of the pipette from the cell with a
concurrent loss of capacitive transient demonstrated that the whole
cell patch had been maintained throughout the experiment.
![]() |
(1) |
Cell-attached patches.
The total current recorded in the cell-attached patch is defined as
![]() |
(2) |
![]() |
(3) |
Cell capacitance.
Determination of capacitance from whole cell currents was described
previously (17). Briefly, a 20-mV voltage step was given through the pipette command potential. Currents were recorded at 1-ms
time intervals for 10 ms after the voltage step and were subtracted
from the stepped current. The transient current was then fit to a
single exponential decay. Capacitance is determined empirically
![]() |
(4) |
Acquisition and analysis. Single-channel data acquisition and analysis were performed as previously described (42, 44). The patch-clamp signals were monitored via an Axopatch-1D amplifier (Axon Instruments, Burlingame, CA) equipped with a TMA-1 interface (Axon Instruments). An analog-to-digital recorder (model VR-10; Instrutech, Mineola, NY) was used to create a digitized recording of the experimental data on videotape for off-line analysis of single-channel recordings. The signal for cell-attached patches was filtered to tape at 50-1,000 Hz. Cell-attached patch data were acquired at a sampling rate of 100-500 Hz for analysis. Whole cell currents were controlled by a microcomputer via an Axopatch-1D amplifier and directly loaded onto a microcomputer at 1 kHz. Data acquisition and analysis were carried out with the pClamp suite of programs (version 6; Axon Instruments, Foster City, CA) and SigmaPlot (version 5.0; SPSS, Chicago, IL).
Solutions. Normal saline solution was (in mM) 105 NaCl, 3 KCl, 2 CaCl2, 1.25 MgSO4, 1.25 KH2PO4, 5 HEPES, and 5 dextrose, pH adjusted to 7.6 with NaOH. Dissecting solution and eversion solution consisted of normal saline solution plus 1 g/100 ml of fraction V bovine serum albumin. The solution filling the cell-attached pipette was normal saline solution.
Whole cell bath solution consisted of (in mM) 105 Na gluconate, 4 KCl, 3 CaCl2, 3 MgCl2, 10 HEPES, and 5 dextrose, pH adjusted to 7.6 with NaOH. In some experiments we included 2 mM BaCl2 in this solution. To keep osmolarity constant, 1 mM CaCl2 and 1 mM MgCl2 were replaced with BaCl2. Whole cell pipette solution consisted of (in mM) 102 K gluconate, 6 KCl, 4 MgCl2, 3 EGTA, 10 HEPES, 5 dextrose, and 2 K2ATP. The pH was adjusted to 7.4 with KOH. When potassium channel inhibitors were included in the whole cell pipette solution, the saline contained (in mM) 76 K gluconate, 6 KCl, 3 MgCl2, 15 D-gluconic acid, 20 tetraethylammonium (TEAOH), 10 CsOH, 3 EGTA, 10 HEPES, 5 dextrose, and 2 K2ATP. The pH was adjusted to 7.4 with HCl. Insulin (bovine) was added to normal bath or whole cell bath solution, resulting in a final concentration of 1 mU/ml. Amiloride (gift from Merck, Sharp, and Dohme) was made up in whole cell bath solution to a concentration of 2 µM. Heptanol was dissolved in pure dimethyl sulfoxide (DMSO). The heptanol-DMSO solution was added directly to the whole cell bath. The apical surface was perfused with either 1 mM heptanol for 3 min or 2 mM heptanol for 5 min. The final concentration of DMSO was 0.25 or 0.5%. The osmolarity of all solutions was monitored and adjusted to 220-232 mosM. All chemicals were purchased from Sigma (St. Louis, MO) unless specified.Statistics. Statistical analysis and graphs were completed with Sigma Plot 2001 (SPSS). Values are expressed as means ± SE. Statistical significance was determined by comparing the difference between untreated tubules and treated tubules with Student's t-test or one-tail test for the difference between independent means. A P < 0.05 was considered statistically significant.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Measurement of GT.
We established that stable values of GT could be
recorded from single cells of the initial collecting tubule. Figure
1A shows representative
current traces in response to a series of 100-ms voltage steps applied
from a holding potential of 80 mV. The voltage steps ranged from
100 to 0 mV in 20-mV increments and were recorded 5 min after whole
cell seal formation. A GT of 7.9 nS for the cell
in Fig. 1A was determined from the slope of the current-voltage plot (Fig. 1B). GT
varied over time after seal formation as illustrated in Fig.
1C. GT was determined immediately after breaking into the cell and was followed for up to 17 min in a
group of six cells. A decline in GT was fit with
an exponential decay function with a time constant of 3 min.
GT was essentially stable by 5 min (8.5 ± 0.7 nS; n = 6) and was not statistically different from
values obtained at 17 min (7.3 ± 1.2 nS; n = 5). The course of initial decay in GT was not
investigated, but it may reflect dialysis with the whole cell pipette
solution contents or some time-dependent increase in the seal
resistance. We chose to determine GT in all
future experiments at least 5 min after seal formation so that all the
solution changes presented to the apical or basolateral membrane
occurred when currents were stable.
|
|
Measurement of GNa.
We used two different approaches in this study to estimate sodium
conductance. In the first, the total and amiloride-insensitive conductances were determined from the slope of the current-voltage plot
as shown in Fig. 2. The amiloride-sensitive change in
GT (GT) was determined
by subtracting the amiloride-insensitive slope conductance from the
GT. Using this method of subtracting the two
slopes assumes that amiloride blocks only the sodium conductance. Figure 3A shows a linear
increase in
GT, with time, after exposure of
the apical membrane to amiloride (2 µM). The mean values of
GT were significantly higher at 5 (P < 0.05) and 8 (P < 0.005) min
after application of amiloride (2 µM) compared with the 1 min value.
Because the difference in slope conductances is not a true
GNa in our model system, we refer to this value
as
GT.
|
Cell coupling. Connexins allow electrical communication to occur between cells and have been reported to be present in the kidney (4, 19). If current passes between cells via connexins, our method of measuring GNa would provide an overestimate. Importantly, efforts to evaluate experimental perturbations that change the GNa could be complicated by any changes in the resistance of the connexin and not the channel under observation. Therefore, to accurately measure single-cell conductances, the cells must be electrically isolated. No information is available regarding the presence of connexins or electrical coupling of principal cells in the collecting tubule of Ambystoma.
One way we addressed this question was by measuring the capacitance of the cell. Theoretically, coupled cells would present multiexponential decay of current, whereas electrically isolated cells would express a single exponential decay transient (13, 17). The measurement of cell capacitance was determined by fitting the transient current response from a +20-mV step to a single exponential decay (Fig. 4). The fit line in Fig. 4 had a time constant of 0.566 ms. In the experiments in which the capacitance could be determined in untreated tubules, the average capacitance was 63 ± 5 pF (n = 6; Table 1). In 15% of the cells studied, the transient decay was too rapid and resulted in a poor fit. Because the line was best fitted to a single exponential, not a multiexponential, the cells were not well coupled, if coupled at all (13).
|
|
|
Effect of insulin. Table 1 compares measurements of whole cell parameters from untreated initial collecting tubules and tubules pretreated with insulin (1 mU/ml) by perfusion of the basolateral membrane for 10-60 min. The mean capacitance in cells was not different in the two groups, indicating that membrane surface area is similar in both groups. After a 1-min application of 2 µM amiloride to the apical membrane, there is an expected hyperpolarization that is significant in both the control and insulin-pretreated tubules (P < 0.01). The normalized GNa is 70% higher in the insulin-pretreated tubules compared with the untreated tubules (2.0 ± 0.4 nS vs. 1.2 ± 0.3 nS; P < 0.05). Thus insulin does increase the sodium conductance and presumably the sodium transport across the apical membrane in the principal cells of the Ambystoma collecting tubule.
Cell-attached patch clamp. To investigate the single-channel mechanism responsible for the increase in the normalized GNa, we used the cell-attached patch-clamp technique on the apical membrane of the everted initial collecting tubule. Three changes in ENaC activity could account for the increase in GNa: 1) an increase in the number of active channels, 2) an increase in open probability, and 3) an increase in conductance.
Others, when studying the effect of vasopressin on ENaC, suggested that the presence of the patch-clamp pipette might have prevented the incorporation of new channels into the membrane under the patch pipette (26). Because this limitation could also apply to our tubules, we chose to evaluate the effect of insulin by comparing control tubules to insulin-treated tubules (1 mU/ml). Seals were formed on the apical membrane of principal cells after the basolateral surface was perfused for 10-60 min with an appropriate saline solution. Table 2 shows the results of the above experiments. There was no statistical difference between the control and insulin-pretreated (1 mU/ml) tubules with regard to single-channel conductance or pipette reversal potential. The data in Table 2 reporting NPo were collected from cell-attached patch-clamp experiments in which the patch could be held for a long period of time (see MATERIALS AND METHODS). The average NPo for insulin-pretreated (1 mU/ml) cells was 2.6 ± 0.9 (n = 6), a value 70% greater than that seen in untreated cells (1.5 ± 0.7, n = 6). The apparent change in NPo was not statistically significant. It is of interest that our whole cell sodium conductance also increased by 70% in insulin-treated cells (see Table 1). The failure to see a significant increase in NPo is presumably related to the fact that there is a wide variability in the number of active levels per patch. However, the open probability of ENaC was significantly higher (P
|
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
There are two cardinal observations of this study. First, we demonstrate the ability to measure ionic conductances in single principal cells of everted Ambystoma initial collecting tubules. Second, this is the only study in native renal tissue to demonstrate an effect of insulin on the amiloride-sensitive ENaC located on the apical membrane.
We previously reported (14, 43-45) that the collecting tubules of both rats and salamanders can be everted and perfused in vitro. One important advantage of the everted tubule as a biological preparation is that it provides access to the apical membrane for electrophysiological studies while permitting independent control of the saline solution to which the apical and basolateral membranes are exposed.
Two types of cells are present in the initial collecting tubule, principal cells and intercalated cells. The principal cells have characteristically been described as polygonal in shape and are involved in sodium transport, whereas the intercalated cells are rounded or elongated in shape and are involved in hydrogen ion transport (18, 29). In rabbit and rat collecting duct, the distribution of the amiloride-sensitive ENaC is limited to principal cells, not intercalated cells.
Initially, we made no attempt to identify the type of cell being
studied. The pooled resting membrane potential of these cells averaged
76 ± 1 mV (n = 18). This is a value very close
to the potassium equilibrium potential (
84 mV) calculated from the
concentrations of potassium in our bath and pipette solutions.
Therefore, the predominant conductance was for potassium, suggesting
viability of the cells studied. The GT of these
same cells was stable and averaged 8.2 ± 1.2 nS
(n = 18)
5 min after seal formation, another observation consistent with healthy cells. This value is similar to
that reported for rat cortical collecting duct principal cells (17) and cultured mouse cortical collecting duct cells
(23).
We used the amiloride sensitivity of GT as a feature to distinguish principal cells from intercalated cells. Nearly 83% (19 of 23 cells) of the cells in which whole cell patches were treated with amiloride had an amiloride-sensitive conductance. This value was comparable to a 75% incidence of principal cells reported in the initial collecting tubule of a similar salamander, Amphiuma means (38). In previous studies that used cell-attached patch-clamp methods, nearly 80% of successful seals expressed ENaC (44, 45).
The whole cell patch-clamp technique allows the investigator to electrically and chemically control the cell. The cells must be electrically isolated for this experimental technique to yield data that represent the functions of a single cell and not that of a syncytium. In a syncytium, gap junction proteins (connexins) establish cytoplasmic continuity between adjacent cells for electrical and metabolic communication (4). Nine different connexins, at the mRNA or protein level, have been identified in the kidney (19). Freeze-fracture electron and scanning microscopy show gap junctions to be present in the proximal tubule segment but not in the distal tubule segment in reptiles (33), amphibians (22, 38) and mammals (24). In the rat cortical collecting duct, the lack of movement of a fluorescent dye into neighboring cells after it was injected into a principal cell suggests that the cells of the mammalian distal nephron are not coupled (17).
We used two approaches to test for the presence of cell coupling in our tissue. First, we determined the capacitance of the principal cells under investigation. Others have noted that the presence of cell coupling should present capacitive discharge profiles that fit a multiple exponential decay (13). The capacitive discharge of our cells fit a single exponential decay, suggesting that the cells were not electrically coupled.
In the rat distal nephron other groups calculated the cell capacitance from known morphometric estimates of surface area of principal cells, which compares favorably to measured capacitance (17). [It is widely accepted that biological membranes have a specific capacitance of 1 µF/cm2 (13).] Although morphometric estimates of cell surface area are lacking for our species, they do exist for Amphiuma (38), a species very similar to Ambystoma. From their values of surface area of the amphibian principal cell, we calculate the capacitance to be 57 pF/cell. Our value of capacitance in untreated tubules is 63 ± 5 pF/cell. The similarity between these values argues that there is little, if any, electrical coupling between principal cells in our preparation.
The second method to test for electrical coupling is to apply an agent known to uncouple connexins, heptanol, while monitoring the current (1). If cells are coupled, then heptanol should decrease the total cell conductance. We observed no change in total cell conductance when heptanol was added to the apical surface. This evidence also supports the notion that the cells of the amphibian initial collecting tubule are not well coupled.
The apparent viability and stability of the everted initial collecting
tubule preparation made it ideal for studying the regulation of
conductance of various ions in individual principal cells. When we
measured the sodium conductance by subtracting the
GT measured in the presence of amiloride from
that in the absence of amiloride, as shown in Fig. 3A, the
resultant measurements of conductance were not stable. A linear
increase in the conductance persisted for at least 8 min after the
application of amiloride. The GT rose from
1.6 ± 0.3 to 3.5 ± 0.5 nS. This observation surprised us
because it is generally accepted that the time course of amiloride
blockage of sodium channels is very rapid and others have observed a
time-dependent decrease in the amiloride-sensitive sodium conductance
(32). The intercept between the control and amiloride
lines (Fig. 2A) should represent the equilibrium potential for sodium. We expected a more positive value for this intercept (approximately +55 mV). In addition, for reasons we cannot explain, we
observed a great deal of variability in the intercept value (data not
shown). However, the intercept value continued to shift in the negative
direction with time after the amiloride application (data not shown).
This shift would be consistent with an amiloride-dependent decrease in
a potassium conductance. Because the value in Fig. 2A is
near 0 mV after exposure of the apical membrane to amiloride for 1 min,
we presume that the secondary decrease in a potassium conductance has
already begun. We believe the contribution of the nonselective cation
channel would be small, with a relative abundance of 0.07 levels/seal
under control conditions (45). Other investigators were
unable to show the effect of 2 µM amiloride on these nonselective
cation channels (45).
A time-dependent reduction in the activity of small-conductance
potassium channels located in the basolateral membrane of the rat
cortical collecting duct when amiloride is applied to the apical
membrane has already been reported by Lu et al. (25). The
decrease in potassium channel activity continued for several minutes
after the application of amiloride (25), a time course similar to the amiloride-induced GT observed in
this study. We were able to abolish the time-dependent increase in the
amiloride-sensitive GT (Fig. 3) when
potassium channel blockers were present. These data are consistent with
the failure of other investigators to observe secondary effects of
amiloride in the presence of potassium channel blockers
(32). The data suggest that the time-dependent increase in
the amiloride-sensitive
GT observed in Fig.
3A was due to a secondary effect of amiloride on a potassium
conductance. It should be noted that, even in the presence of potassium
channel blockers, the equilibrium value for sodium determined from the intercept of the amiloride and control lines was highly variable (data
not shown).
We decided to estimate GNa under conditions in
which there is little or no potassium conductance. By clamping the cell
at 100 mV, the potassium equilibrium potential, there should be little or no current due to potassium. At this voltage, the change in
current flowing across the membrane is due to sodium because there is
no driving force for potassium movement (35). As shown in
Fig. 3A, we obtained the same initial conductance as before, 1.3 nS; however, GNa was now stable with time.
Although subtraction of the amiloride-insensitive conductance from the
total cell conductance provides a reasonable estimate of
GNa, especially if one measures the
amiloride-insensitive conductance within 1 min of applying the drug,
the presence of a time-dependent change in potassium conductance
certainly reduces its accuracy. We believe that estimates of
GNa determined by measuring the
amiloride-inhibitable current with the cell clamped at 100 mV should
yield more accurate values because they are stable with time and do not
appear to be complicated by time-dependent changes in other ionic
currents. We conclude that the best way to measure
GNa is to measure the amiloride-sensitive
current at or near the potassium equilibrium potential.
That a similar value of GNa can be calculated
from our cell-attached patch-clamp data lends support to the notion
that the GNa presented here is valid. From the
data of Stanton et al. (38), we estimate the surface area
of the apical membrane of principal cells of a similar salamander,
Amphiuma means, to be 349 µm2. Given that the
average number of active ENaC levels is 5.8 levels/patch (Table 2) and
the estimate of the surface area of a single patch is 1.4 µm2 (44), the total number of active levels
for the apical membrane should be 1,446 levels/cell. The whole cell
sodium conductance is given by the following equation
![]() |
It is also possible to calculate the rate of sodium transport of an
initial collecting tubule from the single-cell amiloride-sensitive sodium current measured at 100 mV. The normalized sodium current of
120 pA (Table 1), when divided by Faraday's constant, yields a sodium
flow rate of 124 × 10
17
mol · s
1 · cell
1. Using 60 s/min and assuming 300 principal cells per millimeter of tubule length,
we compute a transport rate of 22.4 pmol · mm
1 · min
1. Stoner
(39) reported a measured sodium transport rate for this
tubule segment of 21.2 pmol · mm
1 · min
1. The
agreement between these values suggests that the methods used to
measure GNa are valid.
The second achievement of this study was to characterize the modulation of ENaC after short-term application of insulin in native renal tissue. It was also the first attempt to study the effect of insulin by measuring both whole cell GNa and then the single ENaC activity. In whole cell patch-clamp experiments, normalized GNa was 1.2 ± 0.3 nS in untreated tubules and 2.0 ± 0.4 nS in tubules pretreated with insulin for 10-60 min. Assuming that the increase in normalized GNa reflects a proportional increase in sodium transport, this represented an increase of 70% in sodium reabsorption at the single-cell level. Increases in net sodium transport of similar magnitude have been reported for the effect of insulin on frog skin (9, 36), toad bladder (5), and A6 cells (2, 6, 15, 27, 34).
Using cell attached patch-clamp methodology, we previously characterized, at the single-channel level, the 5-pS ENaC (42, 44). As reported for the rat collecting tubule (17, 30, 31), this was the only sodium channel we observed under control conditions. For a near doubling of the amiloride-sensitive current to occur after exposure of cells to insulin, there had to be an increase in the single-channel conductance, the number of active channels in the patch, the open probability, or a combination of the three. Cell-attached patch-clamp techniques were used to provide more detail regarding which of these changes in ENaC activity were responsible for the increase in sodium reabsorption.
There has been general agreement that insulin does not cause a
significant change in single-channel conductance of ENaC (2, 6,
15, 27). Several investigators, using noise analysis on A6
cells, showed that insulin increased the number of active apical
membrane channels (2, 6, 15). In contrast, Marunaka et al.
(27), using cell-attached patch-clamp techniques on A6 cells, reported an increase in the open probability of ENaC in the
presence of insulin. Our results in native renal tissue confirmed the
observations of Marunaka et al. (27) and showed a
statistically significant (P 0.01) increase in the open
probability of ENaC in the presence of insulin. This increase was of
similar magnitude to that observed for the whole cell sodium
conductance. These observations, together with the lack of any
measurable effect of insulin on the relative abundance of ENaC or
single-channel conductance, suggest that the primary mechanism by which
insulin increases GNa is to increase the
fraction of time individual ENaC stay in the open state.
The physiological significance of the effect of insulin on sodium reabsorption has not been well defined. Some have suggested that the physiological importance of this hormonal effect resides in the need to increase sodium reabsorption to compensate for increased delivery of sodium to the distal nephron (20). Presumably, increased delivery of sodium is due to increases in glomerular filtration rate associated with eating a large meal. There is little experimental evidence to support this hypothesis. Another possibility is the need to secrete potassium associated with food intake (M. Halperin, personal communication). Elevated activity of ENaC would increase transepithelial voltage, enhancing potassium secretion. There is little experimental evidence to support either hypothesis.
On the other hand, many have suggested that the pathophysiological significance of the effect of insulin on sodium transport could be that hyperinsulinemia would lead to inappropriate sodium reabsorption, volume expansion, and hypertension, a common malady associated with people in a hyperinsulinemic state (3, 6, 10). The direct link between hyperinsulinemia and hypertension remains to be established.
![]() |
ACKNOWLEDGEMENTS |
---|
The authors are grateful to Drs. Peter Holohan, Mitchell Halperin, and Edward Solessio for careful reading of this manuscript and many helpful suggestions. Susan Viggiano gave excellent advice both in the planning of protocols and in manuscript preparation, and Susan Sutterer provided precise and meticulous technical assistance.
![]() |
FOOTNOTES |
---|
This work was supported by National Science Foundation Grant IBN9812314 and American Heart Association Predoctoral Fellowship Grant 0110110T.
Address for reprint requests and other correspondence: L. C. Stoner, Dept. of Neuroscience and Physiology, State Univ. of New York Upstate Medical Univ., 766 Irving Ave., Syracuse, New York 13210 (E-mail: ytallini{at}hotmail.com).
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.
June 26, 2002;10.1152/ajpcell.00606.2001
Received 20 December 2001; accepted in final form 12 June 2002.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Anumonwo, JMB,
Wang HA,
Trabka-Janik E,
Dunham B,
Veenstra RD,
Delmar M,
and
Jalife J.
Gap junctional channels in adult mammalian sinus nodal cells: immunolocalization and electrophysiology.
Circ Res
71:
229-239,
1992[Abstract].
2.
Baxendale-Cox, LM,
and
Duncan RL.
Insulin increases sodium (Na+) channel density in A6 epithelia: implications for expression of hypertension.
Biol Res Nurs
1:
20-29,
1999
3.
Bickel, CA,
Verbalis JG,
Knepper MA,
and
Ecelbarger CA.
Increased renal Na-K-ATPase, NCC, and -ENaC abundance in obese Zucker rats.
Am J Physiol Renal Physiol
281:
F639-F648,
2001
4.
Bruzzone, R,
White TW,
and
Goodenough DA.
The cellular internet: on-line with connexins.
Bioessays
18:
709-718,
1996[ISI][Medline].
5.
Blazer-Yost, BL,
Cox M,
and
Furlanetto R.
Insulin and IGF I receptor-mediated Na+ transport in toad urinary bladders.
Am J Physiol Cell Physiol
257:
C612-C620,
1989
6.
Blazer-Yost, BL,
Liu X,
and
Helman SI.
Hormonal regulation of ENaCs: insulin and aldosterone.
Am J Physiol Cell Physiol
274:
C1373-C1379,
1998
7.
Burg, M,
Grantham J,
Abramow M,
and
Orloff J.
Preparation and study of fragments of single rabbit nephrons.
Am J Physiol
210:
1292-1298,
1966.
8.
Canessa, CM,
Schild L,
Buell G,
Thorens B,
Gautschi I,
Horisberger J,
and
Rossier BC.
Amiloride-sensitive epithelial Na+ channel is made of three homologous subunits.
Nature
367:
463-368,
1994[ISI][Medline].
9.
Civan, MM,
Peterson-Yantorno K,
and
O'Brien TG.
Insulin and phorbol ester stimulate conductive Na+ transport through a common pathway.
Proc Natl Acad Sci USA
85:
963-967,
1988[Abstract].
10.
DeFronzo, RA.
The effect of insulin on renal sodium metabolism.
Diabetologia
21:
165-171,
1981[ISI][Medline].
11.
DeFronzo, RA,
Cooke CR,
Andres R,
Falloona GR,
and
Davis PJ.
The effect of insulin on renal handling of sodium, potassium, calcium, and phosphate in man.
J Clin Invest
55:
845-855,
1975[ISI][Medline].
12.
DeFronzo, RA,
Goldberg M,
and
Agus ZS.
The effects of glucose and insulin on renal electrolyte transport.
J Clin Invest
58:
83-90,
1976[ISI][Medline].
13.
DeRoos, AD,
van Zoelen EJ,
and
Theuvenet AP.
Determination of gap junctional intercellular communication by capacitance measurements.
Pflügers Arch
431:
556-563,
1996[ISI][Medline].
14.
Engbretson, BG,
Beyenbach KW,
and
Stoner LC.
The everted renal tubule: a methodology for direct assessment of apical membrane function.
Am J Physiol Renal Fluid Electrolyte Physiol
255:
F1276-F1280,
1988
15.
Erlij, D,
DeSmet P,
and
van Driessche W.
Effect of insulin on area and Na+ channel density of apical membrane of cultured toad kidney cells.
J Physiol
418.3:
533-542,
1994.
16.
Féraille, E,
Marsy S,
Cheval L,
Barlet-Bas C,
Khadouri C,
Favre H,
and
Doucet A.
Sites of antinatriuretic action of insulin along rat nephron.
Am J Physiol Renal Fluid Electrolyte Physiol
263:
F175-F179,
1992
17.
Frindt, G,
Sackin H,
and
Palmer LG.
Whole cell currents in rat cortical collecting tubule: low-Na diet increases amiloride-sensitive conductance.
Am J Physiol Renal Fluid Electrolyte Physiol
258:
F562-F567,
1990
18.
Garty, H,
and
Palmer LG.
Epithelial sodium channels: function, structure, and regulation.
Physiol Rev
77:
359-396,
1997
19.
Guo, R,
Liu L,
and
Barajas L.
RT-PCR study of the distribution of connexin 43 mRNA in the glomerulus and renal tubular segments.
Am J Physiol Regul Integr Comp Physiol
275:
R439-R447,
1998
20.
Gupta, AK,
Clark RV,
and
Kirchner KA.
Effects of insulin on renal sodium excretion.
Hypertension
19, SupplI:
I78-I82,
1992[ISI][Medline].
21.
Hamill, OP,
Marty A,
Neher E,
Sakmann B,
and
Sigworth FJ.
Improved patch-clamp techniques for high-resolution current recording from cells and cell-free membrane patches.
Pflügers Arch
391:
85-100,
1981[ISI][Medline].
22.
Hinton, DE,
Stoner LC,
Burg MB,
and
Trump BF.
Heterogeneity in the distal nephron of the salamander (Ambystoma tigrinum): a correlated structure function study of isolated tubule segments.
Anat Rec
204:
21-32,
1982[ISI][Medline].
23.
Korbmacher, C,
Segal AS,
Fejes-Tóth G,
Giebish G,
and
Boulpaep EL.
Whole-cell currents in single and confluent M-1 mouse cortical collecting duct cells.
J Gen Physiol
102:
761-793,
1993[Abstract].
24.
Kühn, K,
and
Reale E.
Junctional complexes of the tubular cells in the human kidney as revealed with freeze-fracture.
Cell Tissue Res
160:
193-205,
1975[ISI][Medline].
25.
Lu, M,
Giebisch G,
and
Wang W.
Nitric oxide links the apical Na+ transport to the basolateral K+ conductance in the rat cortical collecting duct.
J Gen Physiol
110:
717-726,
1997
26.
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
27.
Marunaka, Y,
Hagiwara N,
and
Tohda H.
Insulin activates single amiloride-blockable Na channels in a distal nephron cell line (A6).
Am J Physiol Renal Fluid Electrolyte Physiol
263:
F392-F400,
1992
28.
O'Neil, RG,
and
Hayhurst RA.
Functional differentiation of cell types of cortical collecting duct.
Am J Physiol Renal Fluid Electrolyte Physiol
248:
F449-F453,
1985
29.
Pácha, J,
Frindt G,
Sackin H,
and
Palmer LG.
Apical maxi K channels in intercalated cells of CCT.
Am J Physiol Renal Fluid Electrolyte Physiol
261:
F696-F705,
1991
30.
Palmer, LG,
and
Frindt G.
Amiloride-sensitive Na channels from the apical membrane of the rat cortical collecting tubule.
Proc Natl Acad Sci USA
83:
2767-2770,
1986[Abstract].
31.
Palmer, LG,
and
Frindt G.
Gating of Na channels in the rat cortical collecting tubule: effects of voltage and membrane stretch.
J Gen Physiol
107:
35-45,
1996[Abstract].
32.
Palmer, LG,
Sackin H,
and
Frindt G.
Regulation of Na+ by luminal Na+ in rat cortical collecting tubule.
J Physiol
509.1:
151-162,
1998
33.
Peek, WD,
Shivers RR,
and
McMillan DB.
Freeze-fracture analysis of junctional complexes in the nephron of the garter snake, Thamnophis sirtalis.
Cell Tissue Res
179:
441-451,
1977[ISI][Medline].
34.
Record, RD,
Johnson M,
Lee S,
and
Blazer-Yost BL.
Aldosterone and insulin stimulate amiloride-sensitive sodium transport in A6 cells by additive mechanisms.
Am J Physiol Cell Physiol
271:
C1070-C1084,
1996.
35.
Satlin, LM,
Sheng S,
Woda CB,
and
Kleyman TR.
Epithelial Na+ channels are regulated by flow.
Am J Physiol Renal Physiol
280:
F1010-F1018,
2001
36.
Schoen, HF,
and
Erlij D.
Insulin action on electrophysiological properties of apical and basolateral membranes of frog skin.
Am J Physiol Cell Physiol
252:
C411-C417,
1987
37.
Scott, WN,
Slatin SL,
Cobb MH,
and
Reich IM.
Insulin-induced alterations in the lactoperoxidase-catalyzed radioiodination of membrane proteins of the toad bladder epithelium.
Endocrinology
109:
1775-1777,
1981[Abstract].
38.
Stanton, B,
Biemesderfer D,
Stetson D,
Kashgarian M,
and
Geibisch G.
Cellular ultrastructure of Amphiuma distal nephron: effects of exposure to potassium.
Am J Physiol Cell Physiol
247:
C204-C216,
1984[Abstract].
39.
Stoner, LC.
Isolated perfused amphibian renal tubules: the diluting segment.
Am J Physiol Renal Fluid Electrolyte Physiol
233:
F438-F444,
1977
40.
Stoner, LC.
The movement of solutes and water across the vertebrate distal nephron.
Renal Physiol
8:
237-48,
1985[ISI][Medline].
41.
Stoner, LC,
Burg MB,
and
Orloff J.
Ion transport in cortical collecting tubule: effect of amiloride.
Am J Physiol
227:
453-459,
1974
42.
Stoner, LC,
Engbretson BG,
Viggiano SC,
Benos DJ,
and
Smith PR.
Amiloride-sensitive apical membrane sodium channels of everted Ambystoma collecting tubule.
J Membr Biol
144:
147-156,
1995[ISI][Medline].
43.
Stoner, LC,
and
Morley GE.
Effect of basolateral or apical hyposmolarity on apical maxi K channels of everted rat collecting tubule.
Am J Physiol Renal Fluid Electrolyte Physiol
268:
F569-F580,
1995
44.
Stoner, LC,
and
Viggiano SC.
Environmental KCl causes an upregulation of apical membrane maxi K and ENaC channels in everted Ambystoma collecting tubule.
J Membr Biol
162:
107-116,
1998[ISI][Medline].
45.
Stoner, LC,
and
Viggiano SC.
Apical nonspecific cation channels in everted collecting tubules of potassium-adapted Ambystoma.
J Membr Biol
177:
109-116,
2000[ISI][Medline].
46.
Ussing, HH,
and
Zerahn K.
Active transport of sodium as the source of electric current in the short-circuited isolated frog skin.
Acta Physiol Scand
12:
110-127,
1951.
|
HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
Visit Other APS Journals Online |