Feedback inhibition of rat amiloride-sensitive epithelial sodium channels expressed in Xenopus laevis oocytes

Whole body sodium homeostasis, and consequently

extracellular fluid volume and blood pressure regulation,

requires tight control of the reabsorption of Na¤ by

epithelial cells. The amiloride-sensitive epithelial sodium

channel (ENaC) located at the apical membrane of epithelial

cells plays a central role in Na¤ reabsorption by the cells of

the distal nephron, the distal colon and the ducts of exocrine

glands (Garty & Palmer, 1997; Rossier, 1997; Horisberger,

1998). The physiological importance of the ENaC has been

demonstrated in human hereditary diseases associated

either with gain-of-function mutations causing Liddle’s

syndrome, a form of salt-sensitive arterial hypertension

(Shimkets et al. 1994), or loss-of-function mutations causing

pseudo-hypoaldosteronism type 1 (Chang et al. 1996).

Despite the rapid expansion of our knowledge of the

structure and function of the ENaC, which is most probably

an heterotetramer, áµâã (Firsov et al. 1998), our under-

standing of the molecular aspects of ENaC regulation is still

fragmentary (Horisberger, 1998). In kidney and colon

epithelia, aldosterone and vasopressin are the major

hormonal regulators of the ENaC (Garty & Palmer, 1997).

Two other well-characterized phenomena, both of which are

intrinsic to the epithelial cell, are known to help limit the

rate of Na¤ entry into the cell: these are ‘self-inhibition’ and

‘feedback inhibition’. However, the mechanisms responsible

for these phenomena are still poorly understood. Self-

inhibition signifies the inhibition of the Na¤ channel by

extracellular sodium; this form of negative regulation has a

fast time course and may be due to a direct interaction

between extracellular Na¤ and a site within the Na¤ channel

itself (Palmer et al. 1998). Feedback inhibition describes an

inhibition of the ENaC that is secondary to an increase in

the intracellular Na¤ concentration ([Na¤]é) (Turnheim,

1991). Feedback inhibition has been reported in numerous

studies of intact epithelia and also at the cellular level

(Garty & Palmer, 1997). Several mechanisms have been

proposed to explain feedback inhibition: it has been

reported to be mediated by a fall in intracellular pH (pHé)

Journal of Physiology (1999), 516.1, pp.31—43 31

Feedback inhibition of rat amiloride-sensitive epithelialsodium channels expressed inXenopus laevis oocytes

Hugues Abriel and Jean-Daniel Horisberger

Institute of Pharmacology and Toxicology, School of Medicine, University of Lausanne,

Switzerland

(Received 15 September 1998; accepted after revision 6 January 1999)

1. Regulation of the amiloride-sensitive epithelial sodium channel (ENaC) is essential for the

control of body sodium homeostasis. The downregulation of the activity of this Na¤ channel

that occurs when the intracellular Na¤ concentration ([Na¤]é) is increased is known as

feedback inhibition. Although intracellular Na¤ is the trigger for this phenomenon, its

cellular and molecular mediators are unknown.

2. We used the ‘cut-open oocyte’ technique to control the composition of the intracellular

milieu of Xenopus oocytes expressing rat ENaCs to enable us to test several factors

potentially involved in feedback inhibition.

3. The effects of perfusion of the intracellular space were demonstrated by an

electromicrographic study and the time course of the intracellular solution exchange was

established by observing the effect of intracellular pH: a decrease from pH 7·4 to 6·5

reduced the amiloride-sensitive current by about 40% within 2 min.

4. Feedback inhibition was observed in non-perfused oocytes when Na¤ entry induced a large

increase in [Na¤]é. Intracellular perfusion prevented feedback regulation even though the

[Na¤]é was allowed to increase to values above 50 mÒ.

5. No effects on the amiloride-sensitive current were observed after changes in the

concentration of Na¤ (from 1 to 50 mÒ), Ca¥ (from 10 to 1000 nÒ) or ATP (from nominally

free to 1 or 5 mÒ) in the intracellular perfusate.

6. We conclude that feedback inhibition requires intracellular factors that can be removed by

intracellular perfusion. Although a rise in [Na¤]é may be the trigger for the feedback

inhibition of the ENaC, this effect is not mediated by a direct effect of Na¤, Ca¥ or ATP on

the ENaC protein.

8737

Keywords:

(Harvey et al. 1988) or a rise in intracellular calcium (Silver

et al. 1993) and to involve G proteins (Gái2 or Gáï) or intra-

cellular chloride (Komwatana et al. 1998). However, no

consensus has yet emerged. For instance, different and

sometimes conflicting results concerning the direct effects of

Na¤ or Ca¥ on the characteristics of ENaC gating and the

role of these ions in feedback inhibition have been reported

by groups using different experimental approaches (Garty &

Palmer, 1997; Benos et al. 1997).

Although the mechanism responsible for signalling an

increasing [Na¤]é to the Na¤ channel is still not yet clear,

recent findings have cast some light on the effector

mechanisms by which the activity of the channel may be

decreased. Firstly, the gain-of-function mutations of ENaCs

associated with Liddle’s syndrome have been shown to

strongly decrease the sensitivity of the ENaC to an [Na¤]é

increase (Kellenberger et al. 1998). These mutations are

located within a short proline-rich segment (PY-motif) of

the cytoplasmic COOH-terminus of the â- and ã-subunits

(Schild et al. 1996). This region interacts with the newly

described cytosolic protein Nedd4 (Staub et al. 1996) which

bears WW-domains known to bind to PY-motifs and a

ubiquitin-protein ligase domain. However, the proposed

mechanism — Nedd4 binding to the PY-motif followed by

ubiquitination of the ENaC and its targeting for degradation

— has yet to be fully demonstrated. Secondly, Shimkets et

al. (1997) showed that overexpression of a dominant-

negative mutant of dynamin in Xenopus oocytes resulted in

an increase in the half-life of wild-type co_expressed ENaCs,

but not of Liddle-mutant channels. These observations

suggest that both ubiquitination and internalization of

ENaCs via clathrin-mediated endocytosis are regulatory

mechanisms that may play an important role in feedback

inhibition, but the relationship between these two

mechanisms is not yet understood.

In this study, we have examined the role of several factors

potentially involved in feedback inhibition using the cloned

rat ENaC (rENaC) expressed in a well-characterized

expression system, the Xenopus oocyte. In order to obtain a

precise and rapid control of the intracellular milieu, we used

the so-called ‘cut-open oocyte’ technique, which permits

intracellular perfusion (Taglialatela et al. 1992). Although

feedback inhibition could be observed in the absence of

intracellular perfusion, it did not occur when the intra-

cellular side of the membrane was efficiently rinsed with

solutions of high or low Na¤ concentration. Under these

conditions, the amiloride-sensitive conductance (GAmil) was

rapidly and reversibly inhibited by intracellular

acidification, but was not influenced by changes in intra-

cellular Na¤, ATP or Ca¥. These results suggest that feedback

inhibition in the Xenopus oocyte, which is triggered by an

increase in [Na¤]é (Kellenberger et al. 1998), is not due to a

direct interaction of Na¤, ATP or Ca¥ with the ENaC and

that, under our experimental conditions, one or more critical

components of the feedback inhibition mechanism are

removed by intracellular perfusion.

METHODSENaC expression inXenopus oocytes

Stage V—VI oocytes were surgically removed from the ovarian

tissue of female Xenopus laevis which had been anaesthetized by

immersion in MS-222 (2 g l¢; Sandoz, Basel, Switzerland).

Following surgery, the frogs were allowed to recover in isolation in

a shallow tank and, after full recovery had been verified a few

hours later, they were returned to the rearing tank. About two

months later, the frogs were operated on a second time for the

removal of the ovarian lobe on the other side. They were then killed

by decapitation under anaesthesia. All procedures were performed

in accordance with local institutional animal welfare guidelines

(State of Vaud, Switzerland). The oocytes were defolliculated as

described previously (Puoti et al. 1995) and were pressure-injected

at the border between the vegetal and animal poles with 50 nl of a

solution containing equal amounts of the áâã cRNAs of the rENaC

subunits (total quantity, 10 ng per oocyte). The site of injection was

chosen so as not to injure the vegetal pole (see below). The

âR564stop mutant (Liddle-mutant) cRNA was generously provided

by L. Schild, Lausanne, Switzerland. After injection, the oocytes

were kept in modified Barth’s solution (MBS) containing 1 mÒ Na¤

to prevent an increase in [Na¤]é and thereby allow observation of

sodium-dependent downregulation. Electrophysiological measure-

ments were performed at room temperature (20—25°C), 14—40 h

after cRNA injection.

Electrophysiological measurements

In this study, we used the cut-open oocyte technique, which was

originally developed by Taglialatela et al. (1992). Briefly, a Xenopus

oocyte was mounted between two compartments with the studied

vegetal pole upwards, as preliminary experiments showed a larger

current at this pole. This correlates well with the much higher

immunocytochemical staining for rENaCs at this pole than at the

animal pole (J. Loffing, personal communication). As shown in

Fig. 1, the superior pole of the oocyte was in contact with the upper

bath through a hole of •500 ìm in diameter. The middle (guard)

bath served to provide electrical isolation between the upper

(extracellular) and lower (intracellular) compartments through

independent voltage clamping of the middle bath at the same

electrical potential as the upper bath. The upper (extracellular)

compartment was superfused by gravity (flow rate, •6 ml min¢)

with an extracellular sodium-containing solution (see below). The

lower pole of the oocyte was impaled with a glass microelectrode

which was simultaneously used as an intracellular perfusion pipette

and a voltage-recording electrode. This modification of the original

set-up was first described by Costa et al. (1994). The resistance of

the electrode, when filled with the intracellular solutions described

below, was about 0·2—0·7 MÙ. For the purpose of intracellular

perfusion, the pipette was advanced into the oocyte until it was

just visible from above through the membrane and yolk. We chose

the flow rate for perfusion so that we could observe a ‘washing-out’

of the yolk platelets, and with time the membrane became

translucent. Only oocytes in which this was observed were

considered to be intracellularly well perfused and only these were

used for further experiment and analysis. In order to obtain this

effect, the flow rate needed to be between 1 and 6 ìl min¢. Higher

rates almost always caused a rapid and marked loss of membrane

resistance or created visible holes in the membrane. The solution

was perfused by means of a precision syringe pump (Infors AG,

Basel, Switzerland). In order to minimize the dead space when the

perfusion solution was changed, we introduced two thin capillaries

by which test solutions were introduced into the perfusion pipette

close to the tip. The remaining dead space was about 2—5 ìl. The

H. Abriel and J.-D. Horisberger J. Physiol. 516.132

voltage clamp was performed using a Dagan cut-open oocyte

voltage-clamp apparatus (Dagan Corporation, Minneapolis, MN,

USA; Model CA-1 High Performance Oocyte Clamp). Data

acquisition and analysis were performed using a TL1 DMA digital

converter system and the pCLAMP software package (Axon

Instruments, Foster City, CA, USA; version 5.5). The holding

potential was −100 mV. Series of 175 ms voltage pulses were

applied to vary the membrane potential from the holding potential

to levels within the range −140 to +60 mV in increments of 20 mV.

The current signal was filtered at 50 Hz using a 4-pole Bessel filter.

The amiloride-sensitive Na¤ current (IAmil) was defined as the

difference between the Na¤ currents obtained with and without

5 ìÒ amiloride (Sigma) in the upper bath. The amiloride-sensitive

conductance (GAmil) was measured between −80 and −100 mV. The

apparent intracellular Na¤ concentration, [Na¤]é, was calculated

from the reversal potential (Erev) of the amiloride-sensitive current

using the following formula:

aNaé RT–––= exp (Erev–– ),aNaï F

where aNaé and aNaï are the intracellular and extracellular Na¤

activities, respectively, F is the Faraday constant, R is the

Boltzmann constant and T is the temperature. As intracellular and

extracellular solutions were of similar ionic strength, we assumed

that the ratio of intracellular to extracellular Na¤ concentrations

was approximately the same as the ratio of intracellular to

extracellular Na¤ activities. We have therefore reported all the results

as apparent Na¤ concentrations. Data are shown as means ± s.e.m.

Solutions and chemicals

After cRNA injection, the oocytes were incubated in a low-Na¤

MBS containing (mÒ): 1 NaCl, 60 N-methyl-ª_glutamine-Cl

(NMDG), 40 KCl, 0·8 MgSOÚ, 0·3 Ca(NO×)µ, 0·4 CaClµ, 10 Hepes-

NMDG (pH 7·2). The extracellular solution for the electro-

physiological experiments had the following composition (mÒ): 0·8

MgSOÚ, 0·4 CaClµ, 5 BaClµ, 10 tetraethylammonium-Cl (TEA), 10

Hepes-NMDG (pH 7·4), the various Na¤ concentrations indicated in

the text being obtained by appropriate addition of sodium

gluconate and NMDG-gluconate to give a total concentration of

100 mÒ. The solution used for the intracellular perfusion contained

(mÒ): 0·8 MgClµ, 1 or 5 ATP, 2 EGTA, 10 Hepes-NMDG, and the

pH was varied from 6·5 to 7·4. The various Na¤ concentrations

indicated in the text (i.e. 1, 20 or 50 mÒ) were obtained by adding

appropriate amounts of sodium gluconate and potassium gluconate

to give a total concentration of 100 mÒ. The various concentrations

of free Ca¥ in the intracellular solution were obtained by adding

appropriate amounts of CaClµ. The free Ca¥ concentrations were

calculated using DOS software, taking into account the pH, ATP

and EGTA concentrations (Calcium v1.1; Chang et al. 1988).

Electron microscopy

The oocytes used for the electron microscopy experiments were the

ones previously used for the electrophysiological measurements. To

Feedback inhibition of epithelial sodium channelJ. Physiol. 516.1 33

Figure 1. Schematic illustration of oocyte intracellular perfusion by the cut-open oocytetechnique

Illustration of an oocyte (•1 mm in diameter) mounted in the cut-open oocyte chamber (not shown). The

chamber consisted of three compartments, of which the upperÏextracellular compartment was continually

perfused and was in contact with the exposed membrane of the oocyte through a hole of •500 ìm

diameter. The guard compartment allowed for electrical isolation between the upper bath and the

lowerÏintracellular bath. For details of the electrical circuit, see Taglialatela et al. (1992) and Costa et al.

(1994). The pipette for perfusion and voltage recording (tip •100 ìm) was inserted into the animal (dark)

pole of the oocyte and advanced to just below (100—300 ìm below) the membrane. The flow of solution

(filled arrows) removed almost all visible yolk platelets below the studied membrane and formed a yolk-free

‘cone’ in the middle of the cell. The solution flowed back and around the pipette and through the opening

made by the impalement.

H. Abriel and J.-D. Horisberger J. Physiol. 516.134

Figure 2. Electron micrograph of the plasma membrane of control and perfused oocytes

A, cortical region of one representative oocyte expressing ENaCs, which was voltage clamped at −100 mV for 20 min

using the two-electrode voltage-clamp technique. The microvilli of the plasma membrane, just below the vitelline

membrane (VM) surrounding the oocyte, can be easily recognized. Yolk platelets (Y) and cortical (C) and pigment (P)

granules can be seen below the membrane. Dense ferritine patches are attached to the vitelline membrane. B, electron

micrograph of the membrane of an oocyte which had been perfused using the cut-open oocyte set-up. The general

architecture of the microvilli was not modified. However, the density of the cytosolic granulations below the

membrane and within the microvilli was clearly decreased. Despite the perfusion, a layer of cytosolic structures (yolk

platelets, cortical and pigment granules) remained attached to the membrane. This micrograph illustrates the three

postulated zones within which the intracellular and extracellular perfusions do not cause convectional flux and where

the Na¤ concentration is influenced only by diffusion: (i) the space between the vitelline and plasma membranes,

(ii) the compartment within the microvilli, and (iii) the layer of remaining cytosolic structures. Scale bars, 10 ìm.

enable us to follow membrane trafficking, in some experiments

ferritine was added to the extracellular solution at 0·7 mg l¢

(Dersch et al. 1991). Immediately after the experiment, the oocytes

were fixed, using 1% glutaraldehyde in the measuring Na¤ solution,

at room temperature for 2 h. Only the dissected vegetal pole of the

oocyte was then used in order to reduce the volume of the sample.

The preparations were washed three times in a phosphate-buffered

solution (PBS; mÒ: 137 NaCl, 2·7 KCl, 1·5 KHµPOÚ, 8·3 NaµHPOÚ)

and were postfixed in 4% OsOÚ for 1 h at room temperature. After

three PBS washes, the samples were dehydrated in ethanol and

embedded in Epon 812. Thin sections (50—80 nm) were then

contrasted with uranyl acetate and lead citrate and observed by

electron microscopy.

RESULTSUltrastructural morphology of the perfused oocyte

For a better understanding of the results of the electro-

physiological measurements performed using the cut-open

oocyte technique, we studied the effects of intracellular

perfusion at the ultrastructural level. Figure 2A shows an

electron micrograph of the cortical region of a representative

oocyte expressing ENaCs, which had been clamped for

20 min at −100 mV using the classical two-electrode

voltage-clamp technique. Figure 2B shows a fragment

(corresponding to the exposed active membrane in Fig. 1) of

an oocyte that had been perfused using the cut-open oocyte

technique and held at a membrane potential of −100 mV for

the same period of time. It is clear that the intracellular

perfusion had removed all intracellular structures except for

those in a 10—60 ìm submembrane zone. In this zone, a few

cytosolic structures (such as small yolk platelets, pigment

and cortical granules) were still present, probably attached

to the membrane-associated cytoskeleton (Fig. 2B). Similar

findings were obtained in three other perfused oocytes.

However, there were no gross changes in the morphology of

the membrane infoldings or in that of the extracellular

vitelline membrane.

Amiloride-sensitive current recorded using the cut-open oocyte technique

After mounting an rENaC-expressing oocyte in the cut-

open oocyte chamber, a first measurement of IAmil was

made immediately after the impalement of the cell with the

perfusion and voltage-recording pipette. The magnitude of

IAmil was within the range 0·1—3 ìA in each preparation.

Taking into account the fact that with the cut-open oocyte

method the current flowing across only a small fraction

(about 10%) of the oocyte membrane is measured (see

Fig. 1), the current density was in the same range as that

Feedback inhibition of epithelial sodium channelJ. Physiol. 516.1 35

Figure 3. Current—voltage curves obtained using the cut-open oocyte technique

A, current recordings obtained from a cut-open oocyte perfused with intracellular and extracellular

solutions containing 50 mÒ Na¤ during a series of 175 ms square voltage pulses ranging from −140 to

+60 mV. The exposed membrane (at the vegetal pole of the oocyte) had a diameter of •500 ìm. B, current

recordings as in A obtained after application of 5 ìÒ amiloride. C, amiloride-sensitive currents (i.e. A − B).

D, current—voltage relationships for the whole-membrane current (0), residual current after application of

amiloride (þ) and amiloride-sensitive current (IAmil, 1). The currents were measured 150 ms after the

beginning of the voltage pulse. Vm, membrane potential.

observed with the two-electrode voltage-clamp technique in

rENaC-expressing oocytes (Kellenberger et al. 1998).

Figure 3 shows the current—voltage relationships obtained

from a perfused oocyte (extracellular Na¤ concentration

([Na¤]ï) = [Na¤]é = 50 mÒ) before and after application of

5 ìÒ amiloride.

Intracellular unstirred layers

It is apparent in Fig. 3D that, despite the similar nominal

Na¤ concentrations on the two sides of the membrane, Erev

was clearly negative, about −10 mV in the example shown,

yielding a calculated apparent [Na¤]é larger than 50 mÒ.

H. Abriel and J.-D. Horisberger J. Physiol. 516.136

Figure 4. Relationship between the apparent intracellular sodium concentration, [Na¤]é, and theamiloride-sensitive conductance (GAmil) of the exposed membrane

The apparent [Na¤]é values were calculated from the reversal potential of the amiloride-sensitive current

using a nominal external Na¤ concentration of 50 mÒ. All these values (n = 10) were measured after a

30 min period during which the membrane potential was maintained at −100 mV and the intracellular side

was continuously perfused with a 50 mÒ Na¤ solution. During this period, GAmil was stable (i.e. no run-

down). The straight line is the linear regression for [Na¤]é versus GAmil and demonstrates a statistically

significant correlation (r  = 0·81, P < 0·001). Note that the intercept on the ordinate is close to 50 mÒ,

which is the nominal Na¤ concentration of the perfused intracellular solution. This relationship indicates the

existence of a compartment just below the membrane, an intracellular unstirred layer, in which [Na¤]é is

influenced by the inflow of Na¤.

Figure 5. Effect of intracellular perfusion: acidification

A, effect on IAmil resulting from acidification of the intra-

cellular perfusion solution from pH 7·4 to 6·5 (filled bars above

the current trace represent application of 5 ìÒ amiloride). At a

flow rate of 5 ìl min¢, acidification decreased IAmil by about

40%; IAmil reached a new steady state after about 2 min.

When the pH was returned to control, in this example, IAmil

reached about 85% of the initial control current. The holding

potential was −100 mV and the downward current deflections

are due to the voltage steps to −60 mV used to monitor the

membrane conductance. B, current—voltage relationships for

IAmil before (0), during (1) and after (þ) a 3 min exposure to a

pH 6·5 intracellular solution. In this example, the pH effect

was fully reversible. The extracellular [Na¤] was 50 mÒ and the

perfused [Na¤] was 1 mÒ.

This discrepancy between the intracellularly perfused Na¤

concentration and the calculated [Na¤]é was observed in

most cases and it was even more obvious when the oocyte

was perfused with a solution containing 1 mÒ Na¤. These

observations indicated that we were not able to control

precisely the Na¤ concentration in the unstirred layers just

below the membrane, even though solution exchange was

effectively taking place within a few tens of micrometres of

the membrane (see Fig. 2B). As the holding potential was

maintained at −100 mV, we hypothesized that a large entry

of Na¤ through the ENaC maintained an increased Na¤

concentration in these unstirred layers. This interpretation

was supported by the finding that the calculated [Na¤]é values

were related to the rate of entry of Na¤ into the cell, as

shown by the linear relationship between apparent [Na¤]é

and measured GAmil (Fig. 4). Indeed, when the rate of entry

of Na¤ through the ENaC was small, the apparent [Na¤]é

corresponded well to the Na¤ concentration in the intra-

cellular perfusate.

Time course of intracellular perfusate exchange andthe effect of pH

The highly Na¤-selective amiloride-sensitive channels are

sensitive to the intracellular pH (Palmer & Frindt, 1987;

Harvey & Ehrenfeld, 1988). In order to test the effectiveness

and the rate of exchange of the intracellular perfusate, we

studied the effect of a change in pHé on IAmil. Reducing the

pH of the perfusate from 7·4 to 6·5 caused a rapid decrease

in IAmil by 46 ± 3% (n = 5). As shown in Fig. 5A, with an

intracellular perfusion flow rate of 5 ìl min¢, IAmil reached

a new steady state after about 2 min. This effect was

usually reversible (Fig. 5A and B): after a return to the

pH 7·4 solution, IAmil recovered to 99 ± 11% of its initial

value.

Feedback inhibition in perfused and non-perfusedoocytes

Kellenberger et al. (1998) observed that when oocytes with

an initially low intracellular Na¤ concentration were

clamped at −100 mV using the two-electrode voltage-clamp

technique, their apparent [Na¤]é increased rapidly and a

large run-down of IAmil was observed. As shown in Fig. 6A,

we observed a similar run-down of IAmil in the cut-open

oocyte setting when the oocytes were not perfused, the

pipette being inserted into the oocyte solely to record the

intracellular electrical potential. As IAmil is influenced by

the reduction in the driving force that results from an

increase in [Na¤]é, we quantified the run-down of the ENaC

by following the calculated GAmil between −80 and −100 mV,

GAmil being almost independent of [Na¤]é at highly negative

Feedback inhibition of epithelial sodium channelJ. Physiol. 516.1 37

Figure 6. Inhibition of ENaC downregulation during intracellular perfusion

Original current recordings under cut-open oocyte conditions showing the decrease in IAmil when the

membrane was clamped at −100 mV (downward deflections are due to voltage pulses to −60 mV). Filled

bars above the current traces represent application of 5 ìÒ amiloride. In A, the oocyte was impaled by the

pipette but not perfused, the pipette being used only to record the intracellular voltage. The extracellular

solution contained 50 mÒ Na¤. B, when the intracellular side of the oocyte was perfused with a solution

containing 20 mÒ Na¤ (extracellular [Na¤], 20 mÒ), IAmil remained stable over a 30—40 min period. C, a

similar abolition of IAmil run-down was also seen when, after an initial 5—10 min perfusion with a 1 mÒ

Na¤ intracellular solution (up to the time indicated by the arrow), the oocyte was perfused with a 50 mÒ

Na¤ solution (extracellular [Na¤], 50 mÒ). No significant effect on IAmil or GAmil was observed following this

increase in the Na¤ concentration of the intracellular perfusate.

membrane potentials. The mean GAmil decreased to

28 ± 5% (n = 6) of its initial control value (Fig. 7A) after

30 min in non-perfused oocytes. This effect was

concomitant with a marked increase in the apparent [Na¤]é,

as shown in Fig. 7B. By contrast, in oocytes perfused with a

20 mÒ intracellular Na¤ concentration (n = 9), we observed

no decline in IAmil or GAmil over a 30 min period (see

example in Fig. 6B) although the apparent [Na¤]é reached

55 ± 10 mÒ after 30 min. Furthermore, we observed no

run-down of GAmil when, after an initial 5—10 min

perfusion with a 1 mÒ Na¤ intracellular solution, the Na¤

concentration was increased to 50 mÒ (Figs 6C and 7A),

even though the intracellular Na¤ concentration increased to

values largely above 50 mÒ (Fig. 7B). Thus, in perfused

oocytes a run-down of GAmil did not occur despite very high

values of apparent [Na¤]é (up to 84 ± 16 mÒ after 40 min

perfusion at a holding potential of −100 mV). However,

when a 50 mÒ Na¤ intracellular solution was used from the

start of the perfusion, we observed a significant initial run-

down of GAmil (data not shown).

Taken together, these results strongly suggest that the intra-

cellular Na¤ concentration, despite being the probable

trigger for the run-down, is not acting directly on the

ENaC. Rather, feedback inhibition may be due to a cascade

of intracellular events involving other factors that may have

been removed by intracellular perfusion. Possible mediators

of the feedback inhibition that could act directly on the

ENaC include ATP, calcium and pH, and we took advantage

of the cut-open oocyte technique to investigate the effect of

these factors on ENaC activity.

Effect of intracellular ATP

Intracellular ATP and the ratio ATPÏADP are known to

regulate various ion channels and transporters (Hilgemann,

1997). In addition, an increase in Na¤ entry into the cell

may lead to a decrease in [ATP], since Na¤ stimulates the

consumption of ATP by the sodium pump. ATP is therefore

a possible mediator of Na¤-dependent ENaC inhibition. As

described in the Methods section, the intracellular perfusion

solutions contained 1 mÒ ATP (a condition we had chosen

initially in case ATP was a necessary co-factor for the actual

regulatory mechanism). In another set of experiments, ATP

was not included in the intracellular perfusate used at the

beginning of the experiment so that we could test the effect

of addition of ATP on IAmil. Increasing the concentration of

ATP in the solution from nominally zero ATP to 1 or 5 mÒ

did not induce any detectable change in IAmil (Fig. 8A).

Effect of intracellular calcium

Intracellular Ca¥ has been proposed as a mediator of

feedback inhibition in several experimental situations (Silver

et al. 1993; Ishikawa et al. 1998). To test the direct effect of

Ca¥ on the ENaC, we started the intracellular perfusion

with a nominally Ca¥-free solution containing 2 mÒ EGTA

and 1 mÒ ATP, and then added calcium to increase the

calculated free Ca¥ concentration to 1000 nÒ for a 10 min

period. This large increase in the intracellular Ca¥

concentration had no significant effect on IAmil (Fig. 8B).

Role of intracellular pH

We have already described the direct effect of pHé on IAmil

(see above and Fig. 5). As a result of the presence of the

H. Abriel and J.-D. Horisberger J. Physiol. 516.138

Figure 7. Run-down of the amiloride-sensitiveconductance (with a concomitant increase in [Na¤]é) andits inhibition by intracellular perfusion

A, when the oocytes were not perfused (0; [Na¤]ï = 50 mÒ),

GAmil decreased to about 30% of its control value after

30 min of voltage clamping at −100 mV. By contrast, when

the cell was perfused (1) with a 50 mÒ Na¤ intracellular

solution, GAmil remained stable for at least 40 min. For non-

perfused and perfused oocytes, the initial current values

were 1·57 ± 0·31 ìA (n = 6) and 1·78 ± 0·65 ìA (n = 6),

respectively, and the initial conductances were 18·7 ± 7·4

and 16·8 ± 3·5 ìS. For the perfused oocytes, note that the

first GAmil value was measured when the cell was perfused

with 1 mÒ Na¤; the Na¤ concentration was changed to

50 mÒ at the time indicated by the arrow. This change from

a 1 to a 50 mÒ Na¤ intracellular perfusion did not lead to a

modification in GAmil. B, in non-perfused oocytes (0), even

though the oocytes were incubated in 1 mÒ Na¤, the values

of the apparent [Na¤]é had already reached 80 mÒ at the

time of the first measurement. The apparent [Na¤]é then

increased to a mean value of about 200 mÒ. When the

oocyte was intracellularly perfused with 50 mÒ Na¤ (1; first

5 min with 1 mÒ: arrow; see A), the mean apparent [Na¤]é

reached a plateau at about 80 mÒ. The actual values were

between 59 and 128 mÒ after 40 min and were a function of

the measured GAmil (see Fig. 4).

Na¤—H¤ exchanger, an increase in [Na¤]é could result in

intracellular acidification, which in turn could be directly

responsible for the inhibition of the ENaC and thus explain

feedback inhibition (Palmer & Frindt, 1987; Harvey et al.

1988). In order to test this possibility, we studied the pH

sensitivity of a mutant ENaC (the Liddle-mutant,

áâR564stopã), a channel which failed to show down-

regulation in oocytes under similar experimental conditions

(Kellenberger et al. 1998). Acidification of the intracellular

perfusate from pH 7·4 to 6·5 decreased IAmil to 70 ± 4%

(n = 3) of the initial value for this mutant. This observation

suggests that pHé is unlikely to be either the single or main

mediator of feedback inhibition.

DISCUSSIONIn the present study, we looked for intracellular factors that

might be involved in the feedback inhibition of ENaCs

expressed in Xenopus oocytes. The main finding was that

this downregulation did not occur when the cytosol of the

cell was largely removed by intracellular perfusion, even

when the intracellular concentration of Na¤ reached values

as high as those at which feedback inhibition was observed

in non-perfused oocytes. This observation suggests that at

least one essential cytosolic component was removed by the

intracellular perfusion. We subsequently found that none of

the factors Na¤, ATP or Ca¥ had a direct effect on ENaC

activity. Further, although pHé did modulate ENaC activity,

it did not seem to be the main or sole mediator of the

observed downregulation.

Study of ENaCs with the cut-open oocyte technique

The cut-open oocyte technique was originally developed for

the study of gating currents in potassium channels, as this

technique allows for rapid voltage clamping (Taglialatela et

al. 1992). This technique also permits intracellular perfusion

of the oocyte and we took advantage of this to study the

influence of intracellular factors on rENaCs. In perfused

oocytes, we measured IAmil, which was similar in terms of

both magnitude and current—voltage relationship to IAmil

measured using the classical two-electrode voltage-clamp

method. Depending on the oocyte batch, these currents

remained very stable for over 30—40 min (see below). We

tested the effectiveness and the rate of exchange of the

intracellular perfusate by observing the effect of a pH

change on the amiloride-sensitive current. In fact, IAmil

rapidly and reversibly decreased by about 40% when the

pHé was changed from 7·4 to 6·5. Two previous studies have

quantified the dependence of the ENaC on pHé. In frog skin

(Harvey & Ehrenfeld, 1988), GAmil decreased by about 80%

when the pHé was lowered from 7·4 to 6·5. In patch-clamp

experiments on the cells of rat cortical collecting ducts

(Palmer & Frindt, 1987), the open probability (Pï) decreased

by 88% in response to an identical decrease in pHé. One

reason for the smaller change in IAmil in our experiments

could be that we could not control with any precision the pH

just below the membrane because of the unstirred-

compartment phenomenon (see below).

The membrane preparation obtained using the cut-open

oocyte technique brings many advantages to the study of

the action of potential intracellular regulatory factors on

Feedback inhibition of epithelial sodium channelJ. Physiol. 516.1 39

Figure 8. Effect of acute change in intracellular ATPand Ca¥ concentrations

A, ATP at 1 or 5 mÒ did not influence IAmil (absolute initial

values for 1 and 5 mÒ ATP, respectively: 0·4 ± 0·05 ìA,

n = 3 and 1·0 ± 0·4 ìA, n = 3). B, when the calculated free

Ca¥ concentration was increased from less than 10 nÒ to

1000 nÒ, no change in IAmilwas observed (absolute initial

value: 1·2 ± 0·7 ìA, n = 3). Note: in both cases, the nominal

extracellular Na¤ concentration was 90 mÒ and the intra-

cellular Na¤ concentration was 20 mÒ. IAmil values for Ca¥ or

ATP (normalized with respect to Control) were obtained

4—5 min or 3 min, respectively, after the intracellular

solution exchange, the flow rate being 5 ìl min¢ in each case.

membrane transport proteins. The elements of the cytosol

that are not attached to either the membrane or the

membrane-associated cytoskeleton can be removed, and the

concentration of any soluble factor(s) can be controlled by

the intracellular perfusion with a time course of a few

minutes. The difficulty of precisely controlling the sub-

membrane concentration of ions because of the existence of

unstirred layers (see below) can be avoided by working

under conditions in which the rate of net transport is low.

The preparation allows the observation of stable channel

activity for periods of up to 40 min under precisely

controlled conditions. The intracellular perfusion can be

carried out using a small volume of solution (down to

100 ìl) allowing the testing of substances available only in

restricted amounts. Furthermore, for the analysis of ENaC

regulation, it is an advantage that the size of the active

membrane surface is large enough to measure ‘macroscopic’

currents, since the regulation of the activity of this channel

is made difficult in studies of single-channel currents by the

large spontaneous variability seen in the Pï of individual

Na¤ channels (Garty & Palmer, 1997).

Unstirred layers

When the oocyte was not perfused, the apparent [Na¤]é

values, calculated using the Erev obtained from the IAmil

current—voltage curve, were surprisingly high (up to

200 mÒ). A similar observation has already been reported

and discussed by Kellenberger et al. (1998). It should be

pointed out that the values calculated from Erev reflect the

[Na¤]é in a cytosolic compartment close to the plasma

membrane, and not the bulk Na¤ concentration inside the

cell. When the cell was perfused, we found that the

calculated apparent [Na¤]é was usually higher than the

perfused Na¤ concentration and that it was linearly related

to GAmil in the same preparation. We interpret these

observations as indicating that the entry of Na¤ through the

ENaC influences [Na¤]é in the unstirred compartment close

to the membrane. As a matter of fact, when GAmil was small,

[Na¤]é corresponded well to the Na¤ concentration of the

perfusate (Fig. 4), which indicates that although this

compartment could not be reached through convectional

flow, it was nevertheless in diffusion equilibrium with the

bulk of the perfused solution. When interpreting these

observations, it is important to note that the calculation

from the Nernst equation using the measured amiloride-

sensitive Erev (see Methods) yields a measure of the

[Na¤]éÏ[Na¤]ï ratio. We have used the nominal [Na¤]ï in our

calculations of [Na¤]é but the presence of extracellular

unstirred layers (between the vitelline layer and the plasma

membrane, see Fig. 2) could also influence the real value of

[Na¤]ï close to the membrane. It follows that we probably

somewhat overestimated the real [Na¤]ï when using the

nominal extracellular Na¤ concentration for the calculations.

The electron micrographs of the perfused oocytes clearly

show a 10—60 ìm zone in which organelles are present

(Fig. 2B). They presumably remain attached to the membrane

by cytoskeletal elements that have not been removed by the

intracellular perfusion. This zone constitutes an intracellular

unstirred layer. Furthermore, the cytosol within the

microvilli may also represent an unstirred compartment

that is important if the Na¤ channels are expressed on the

villi. Thus the Na¤ concentrations close to the membrane on

both sides may be quite different from the concentrations of

the bulk solutions perfused around and inside the oocyte

when a large flux of Na¤ is moving across the membrane.

Our demonstration of the presence of unstirred layers

points to a possibly physiologically relevant sodium

microdomain within the cell, as also observed for Na¤ in

cardiac cells (Carmeliet, 1992; Wendt-Gallitelli et al. 1993).

The cut-open oocyte technique, which allows perfusion of

both sides of the membrane with solutions of known

composition and a precise determination of the trans-

membrane ion gradient from measurement of the reversal

potential, enables us to characterize in a quantitative way

the effect of the unstirred layers. The sodium concentration

in this compartment may reach values very different from

the whole-cell sodium concentration. This implies that

[Na¤]é measurements made using, for example, intracellular

ion-specific microelectrodes, Na¤-specific dyes or tissue

homogenization need to be interpreted with caution when

the aim is to establish the ion concentration at the plasma

membrane and its effect on membrane proteins such as ion

channels, coupled transport systems or pumps (Carmeliet,

1992; Fujioka et al. 1998).

Feedback inhibition in perfused and non-perfusedoocytes

Feedback inhibition of ENaCs was first proposed by

MacRobbie & Ussing (1961) who observed that cells of the

frog skin expressing apical amiloride-sensitive Na¤ channels

did not swell as expected when Na¤ extrusion by Na¤,K¤-

ATPase was blocked. The presence of a feedback regulation

serving to inhibit the channel when Na¤ enters the cell has

since been confirmed at the single-channel level (Silver et al.

1993; Frindt et al. 1993). In oocytes expressing rENaCs

studied using the two-electrode voltage-clamp technique, a

rapid run-down is consistently observed. This phenomenon

was investigated in detail in a recent study (Kellenberger et

al. 1998) in which it was convincingly shown that the rate of

this run-down was directly dependent on the rate of Na¤

entry into the cell (the higher the IAmil, the faster the rate of

run-down). This indicated that the increase in [Na¤]é is the

trigger for the feedback inhibition of the ENaC expressed in

the Xenopus oocyte. In our experimental setting, it was also

possible to see an increase in the apparent [Na¤]é and a

concomitant run-down of GAmil when the cell was simply

impaled with the voltage-recording pipette (and not

perfused). This run-down was of the same magnitude (by

about 70—80% in 30 min) as that seen in two-electrode

voltage-clamp experiments (Kellenberger et al. 1998). The

higher values of apparent [Na¤]é in our study (an average of

187 versus 120 mÒ) may be explained by the fact that we

were able to use oocytes with a larger IAmil (about 15—20 ìA

when extrapolated to the whole oocyte as compared with

H. Abriel and J.-D. Horisberger J. Physiol. 516.140

4—8 ìA per oocyte in the study of Kellenberger et al. 1998).

In the cut-open oocyte setting we are not limited by very

large inflows of Na¤ since osmotic swelling of the oocyte

cannot occur.

Although feedback inhibition was clearly observed in non-

perfused oocytes, the Na¤-dependent downregulation was

abolished when the oocytes were intracellularly perfused

with various solutions, even though the [Na¤]é reached high

values (within the range 50—130 mÒ). In addition, a change

in the perfused intracellular Na¤ concentration from 1 to

50 mÒ did not induce any detectable change in the

amiloride-sensitive Na¤ conductance. These observations

strongly suggest that sodium itself does not regulate the

channel by direct interaction with the ENaC protein.

Palmer and co-workers (Palmer et al. 1989) reached the

same conclusion after studying Na¤ channel activity in

excised patches from rat cortical collecting ducts. However,

this finding is in conflict with the results of a recent study

involving excised patch-clamp experiments on rENaCs

expressed in MDCK cells (Ishikawa et al. 1998). In that

study, the NPï (mean number of open channels) was

decreased by about 75% when [Na¤]é was increased from 0

to 100 mÒ. We have no documented explanation for this

difference but it may be that, with the inside-out patch-

clamp configuration used in the experiments on MDCK cells,

some essential component of the feedback mechanism

remains associated with the membrane, while it was

removed by intracellular perfusion in the oocytes used here.

The other studies of the effect of intracellular Na¤ on

amiloride-sensitive channels are difficult to compare with

ours because of the different experimental conditions used:

either the channel was studied in an artificial membrane

consisting of a lipid bilayer (Ismailov et al. 1995) or the

work was carried out using whole-cell patch clamping,

which does not allow efficient removal of intracellular

components of large molecular size (Komwatana et al. 1996).

We think that, by using intracellular perfusion of the

oocyte, we were able to remove most of the cytosol,

including even those slowly diffusible cytosolic components

that could be associated with ENaCs such as Nedd4,

ubiquitin, elements involved in clathrin-mediated endocytosis

andÏor other factors like G proteins (as recently proposed in

a study on salivary duct cells; Komwatana et al. 1998).

Thus, our preparation may be devoid of most of the

potential regulatory elements that are neither membrane

proteins nor proteins strongly linked with the membrane-

associated cytoskeleton.

Mediators of feedback inhibition

Several mediators have been implicated in ENaC feedback

inhibition (for a review, see Garty & Palmer, 1997). These

factors may form part of a molecular cascade starting with a

sodium-sensing mechanism and ending with an effector

acting directly on the channel itself. As discussed in detail

above, we have shown that Na¤ is not the effector of this

downregulating mechanism.

ATP was the first regulatory factor to be suggested. ATP is

a known regulator of a whole class of K¤ channels (Tucker &

Ashcroft, 1998) and a change in [Na¤]é modulates the ATP

content of the cell, an increase in [Na¤]é increasing the

consumption of ATP by the cell’s Na¤,K¤-ATPase (Tsuchiya

et al. 1992). However, we failed to observe any effect of ATP

on the amiloride-sensitive Na¤ current when the perfusate

was changed from a nominally ATP-free solution to solution

containing 1 or 5 mÒ ATP. In addition, the run-down of

ENaC activity observed in the absence of Na¤,K¤-ATPase

activity (our present study with a K¤-free extracellular

solution) was similar to that observed when the Na¤,K¤-

ATPase was activated by the presence of extracellular K¤

(Kellenberger et al. 1998). Thus our results do not support

the hypothesis of a regulation by intracellular ATP and a

direct link between ENaC activity and Na¤,K¤-ATPase

activity.

We next addressed the question of the role of intracellular

Ca¥, a factor proposed a long time ago as a possible

mediator of feedback inhibition (Grinstein & Erlij, 1978;

Schultz, 1981). An increase in [Na¤]é might decrease the

driving force available to the Na¤—Ca¥ exchanger and so

lead to an increase in [Ca¥]é. A number of studies — on intact

epithelia (Ling & Eaton, 1989), MDCK epithelia (whole-cell

patch-clamp experiments; Ishikawa et al. 1998) and

membrane vesicles from toad urinary bladder (Garty et al.

1987) — have demonstrated an inhibitory effect of intra-

cellular Ca¥ on Na¤ conductance. However, under our

experimental conditions, a change from less than 10 nÒ to

1000 nÒ Ca¥ in the intracellular perfusate had no effect on

the ENaC. These negative results with intracellularly

perfused oocytes suggest that the above effects may not be

due to a direct interaction of Ca¥ with the ENaC protein.

Several other studies have indicated that the ENaC

expressed in Xenopus oocytes is not sensitive to [Ca¥]é,

arguing against a direct effect of Ca¥ on the ENaC. The

run-down phenomenon observed by Kellenberger et al.

(1998) was not decreased by buffering [Ca¥]é with EGTA or

BAPTA. Another piece of indirect evidence suggesting the

absence of an inhibitory effect of [Ca¥]é on the ENaC

expressed in oocytes is provided by the observation that

extracellular trypsin, which produces a large increase in

intracellular Ca¥, activates, rather than inhibits, the

amiloride-sensitive Na¤ current (Chra� úbi et al. 1998).

As in the case of intracellular Na¤, the results with Ca¥

indicate that some regulatory components normally present

in epithelial cells are needed for Ca¥ to exert its effect on

ENaC activity. This hypothesis is strongly supported by the

results of Palmer & Frindt (1987), who worked on the cells

of rat collecting ducts. They showed that the single

epithelial Na¤ channel currents observed in excised inside-

out patches were not influenced by [Ca¥]é, while an increase

in [Ca¥]é brought about by the addition of a calcium

ionophore decreased channel activity in cell-attached patch-

clamp recordings.

Feedback inhibition of epithelial sodium channelJ. Physiol. 516.1 41

Intracellular pH has also been proposed as a mediator of

feedback inhibition. Harvey et al. (1988), using the same

type of argument as that used in favour of Ca¥, suggested

that, because of the presence of the Na¤—H¤ exchanger, an

increase in [Na¤]é would result in intracellular acidification

which would then inhibit the Na¤ channel. Like other

workers (Palmer & Frindt, 1987; Harvey & Thomas, 1987;

Harvey et al. 1988), we did indeed observe a significant

inhibition of ENaC activity by low pH. However, two

observations argue against this way of explaining feedback

inhibition. First, a decrease in pHé to 6·5 reduced IAmil by

about 40%; this means that a much greater degree of

acidification would be necessary to reach the 70—80%

decrease in IAmil seen during Na¤ entry. Such a large decrease

in pHé would seem very unlikely to occur under physiological

conditions. Second, we studied the pH sensitivity of ENaCs

carrying Liddle’s mutation (áâR564stopã) which have been

shown to be resistant to Na¤-dependent downregulation

(Kellenberger et al. 1998). Incorporation of this mutation

did not abolish the sensitivity of the ENaC to the intra-

cellular pH and we therefore conclude that protons are most

probably not the effectors for feedback inhibition.

Having excluded a role for intracellular pH and the direct

effects of ATP, Ca¥ and Na¤ itself, what are we left with as

a possible mechanism for feedback inhibition?

Kellenberger et al. (1998) demonstrated that this Na¤-

triggered regulatory phenomenon was dependent on the

presence of an intact PY-motif (Staub & Rotin, 1997) in the

â- and ã-subunits. The cytosolic protein Nedd4 (Staub et al.

1996), which is found in almost all tissues expressing ENaCs

(Staub et al. 1997) and also in the Xenopus oocyte (Staub et

al. 1996), binds to these PY-motifs via its WW-domains.

This interaction probably downregulates the ENaC but the

mechanism underlying this effect has not yet been elucidated.

In another model (viz. mouse salivary duct cells exhibiting

amiloride-sensitive currents), Nedd4 antibodies were shown

to disrupt the Na¤-dependent negative regulation of these

currents (Dinudom et al. 1998). ENaC Na¤-dependent

downregulation (over a period of hours) has recently been

shown to be prevented by the expression of a dominant-

negative mutant of dynamin in Xenopus oocytes (Shimkets

et al. 1997), pointing to a role for clathrin-mediated

endocytosis in this regulation.

Our results show that intracellular Na¤, Ca¥ and ATP do

not interact directly with the ENaC protein expressed in

oocytes. These observations do not mean that these factors

are not involved at all in the regulation of ENaC activity,

rather that the mechanism underlying this regulation

requires the presence of additional factors that can be

removed by intracellular perfusion. Use of the cut-open

oocyte technique should allow the effects of several more of

these factors to be tested.

Benos, D. J., Fuller, C. M., Shlyonsky, V. G., Berdiev, B. K. &

Ismailov, I. I. (1997). Amiloride-sensitive Na¤ channels — insights

and outlooks. News in Physiological Sciences 12, 55—61.

Carmeliet, E. (1992). A fuzzy subsarcolemmal space for intracellular

Na¤ in cardiac cells? Cardiovascular Research 26, 433—442.

Chang, D., Hsieh, P. S. & Dawson, D. C. (1988). Calcium: a program

in BASIC for calculating the composition of solutions with specified

free concentrations of calcium, magnesium and other divalent

cations. Computers in Biology & Medicine 18, 351—366.

Chang, S. S., Grunder, S., Hanukoglu, A., Rosler, A., Mathew,

P. M., Hanukoglu, I., Schild, L., Lu, Y., Shimkets, R. A.,

Nelson-Williams, C., Rossier, B. C. & Lifton, R. P. (1996).

Mutations in subunits of the epithelial sodium channel cause salt

wasting with hyperkalaemic acidosis, pseudohypoaldosteronism

type 1. Nature Genetics 12, 248—253.

Chra� úbi, A., Vallet, V., Firsov, D., Kharoubi-Hess, S. &

Horisberger, J.-D. (1998). Protease modulation of the activity of

the epithelial sodium channel expressed in Xenopus oocytes.

Journal of General Physiology 111, 127—138.

Costa, A. C. S., Patrick, J. W. & Dani, J. A. (1994). Improved

technique for studying ion channels expressed in Xenopus oocytes,including fast superfusion. Biophysical Journal 67, 395—401.

Dersch, M. A., Bement, W. M., Larabell, C. A., Mecca, M. D. &

Capco, D. G. (1991). Cortical membrane-trafficking during the

meiotic resumption of Xenopus laevis oocytes. Cell and TissueResearch 263, 375—383.

Dinudom, A., Harvey, K. F., Komwatana, P., Young, J. A., Kumar,

S. & Cook, D. I. (1998). Nedd4 mediates control of an epithelial Na¤

channel in salivary duct cells by cytosolic Na¤. Proceedings of theNational Academy of Sciences of the USA 95, 7169—7173.

Firsov, D., Gautschi, I., Merillat, A. M., Rossier, B. C. & Schild,

L. (1998). The heterotetrameric architecture of the epithelial sodium

channel (ENaC). EMBO Journal 17, 344—352.

Frindt, G., Silver, R. B., Windhager, E. E. & Palmer, L. G.

(1993). Feedback regulation of Na channels in rat CCT. II. Effects of

inhibition of Na entry. American Journal of Physiology 264,F565—574.

Fujioka, Y., Matsuoka, S., Ban, T. & Noma, A. (1998). Interaction

of the Na¤—K¤ pump and Na¤—Ca¥ exchange via [Na¤]é in a

restricted space of guinea-pig ventricular cells. Journal ofPhysiology 509, 457—470.

Garty, H., Asher, C. & Yeger, O. (1987). Direct inhibition of

epithelial Na¤ channels by a pH-dependent interaction with calcium,

and by other divalent ions. Journal of Membrane Biology 95,151—162.

Garty, H. & Palmer, L. G. (1997). Epithelial sodium channels —

function, structure, and regulation. Physiological Reviews 77,359—396.

Grinstein, S. & Erlij, D. (1978). Intracellular calcium and the

regulation of sodium transport in the frog skin. Proceedings of theRoyal Society B 202, 353—360.

Harvey, B. J. & Ehrenfeld, J. (1988). Role of Na¤ÏH¤ exchange in

the control of intracellular pH and cell membrane conductances in

frog skin epithelium. Journal of General Physiology 92, 793—810.

Harvey, B. J. & Thomas, R. C. (1987). Intracellular pH and calcium

effects on sodium conductance and transport in isolated frog skin

epithelium. Journal of Physiology 394, 92P.

Harvey, B. J., Thomas, S. R. & Ehrenfeld, J. (1988). Intracellular

pH controls cell membrane Na¤ and K¤ conductances and transport

in frog skin epithelium. Journal of General Physiology 92,767—791.

H. Abriel and J.-D. Horisberger J. Physiol. 516.142

Hilgemann, D. W. (1997). Cytoplasmic ATP-dependent regulation of

ion transporters and channels: mechanisms and messengers. AnnualReview of Physiology 59, 193—220.

Horisberger, J.-D. (1998). Amiloride-sensitive Na channels. CurrentOpinion in Cell Biology 10, 443—449.

Ishikawa, T., Marunaka, Y. & Rotin, D. (1998). Electro-

physiological characterization of the rat epithelial Na¤ channel

(rENaC) expressed in MDCK cells — effects of Na¤ and Ca¥. Journalof General Physiology 111, 825—846.

Ismailov, I. I., Berdiev, B. K. & Benos, D. J. (1995). Regulation by

Na¤ and Ca¥ of renal epithelial Na¤ channels reconstituted into

planar lipid bilayers. Journal of General Physiology 106, 445—466.

Kellenberger, S., Gautschi, I., Rossier, B. C. & Schild, L. (1998).

Mutations causing Liddle-syndrome reduce sodium-dependent

downregulation of the epithelial sodium channel in the Xenopusoocyte expression system. Journal of Clinical Investigation 101,2741—2750.

Komwatana, P., Dinudom, A., Young, J. A. & Cook, D. I. (1996).

Cytosolic Na¤ controls an epithelial Na¤ channel via the G(o)

guanine nucleotide-binding regulatory protein. Proceedings of theNational Academy of Sciences of the USA 93, 8107—8111.

Komwatana, P., Dinudom, A., Young, J. A. & Cook, D. I. (1998).

Activators of epithelial Na¤ channels inhibit cytosolic feedback

control — evidence for the existence of a G protein-coupled receptor

for cytosolic Na¤. Journal of Membrane Biology 162, 225—232.

Ling, B. N. & Eaton, D. C. (1989). Effects of luminal Na¤ on single

Na¤ channels in A6 cells, a regulatory role for protein kinase C.

American Journal of Physiology 256, F1094—1103.

MacRobbie, E. A. C. & Ussing, H. H. (1961). Osmotic behaviour of

the epithelial cells of frog skin. Acta Physiologica Scandinavica 53,348—365.

Palmer, L. G. & Frindt, G. (1987). Effects of cell Ca and pH on Na

channels from rat cortical collecting tubule. American Journal ofPhysiology 253, F333—339.

Palmer, L. G., Frindt, G., Silver, R. B. & Strieter, J. (1989).

Feed-back regulation of epithelial sodium channels. Current Topicsin Membranes and Transport 34, 45—60.

Palmer, L. G., Sackin, H. & Frindt, G. (1998). Regulation of Na¤

channels by luminal Na¤ in rat cortical collecting tubule. Journal ofPhysiology 509, 151—162.

Puoti, A., May, A., Canessa, C. M., Horisberger, J.-D., Schild, L.

& Rossier, B. C. (1995). The highly selective low conductance

epithelial Na channel of Xenopus laevis A6 kidney cells. AmericanJournal of Physiology 38, C188—197.

Rossier, B. C. (1997). Cum grano salis — the epithelial sodium channel

and the control of blood pressure. Journal of the American Societyof Nephrology 8, 980—992.

Schild, L., Lu, Y., Gautschi, I., Schneeberger, E., Lifton, R. P. &

Rossier, B. C. (1996). Identification of a PY motif in the epithelial

Na channel subunits as a target sequence for mutations causing

channel activation found in Liddle syndrome. EMBO Journal 15,2381—2387.

Schultz, S. G. (1981). Homocellular regulatory mechanisms in

sodium-transporting epithelia: avoidance of extinction by ‘flush-

through’. American Journal of Physiology 241, F579—590.

Shimkets, R. A., Lifton, R. P. & Canessa, C. M. (1997). The activity

of the epithelial sodium channel is regulated by clathrin-mediated

endocytosis. Journal of Biological Chemistry 272, 25537—25541.

Shimkets, R. A., Warnock, D. G., Bositis, C. M., Nelson-

Williams, C., Hansson, J. H., Schambelan, M., Gill, J. R. Jr,

Ulick, S., Milora, R. V., Findling, J. W., Canessa, C. M.,

Rossier, B. C. & Lifton, R. P. (1994). Liddle’s syndrome: heritable

human hypertension caused by mutations in the beta subunit of the

epithelial sodium channel. Cell 79, 407—414.

Silver, R. B., Frindt, G., Windhager, E. E. & Palmer, L. G.

(1993). Feedback regulation of Na channels in rat CCT. I. Effects of

inhibition of Na pump. American Journal of Physiology 264,F557—564.

Staub, O., Dho, S., Henry, P. C., Correa, J., Ishikawa, T.,

McGlade, J. & Rotin, D. (1996). WW domains of Nedd4 bind to

the proline-rich PY motifs in the epithelial Na¤ channel deleted in

Liddle’s syndrome. EMBO Journal 15, 2371—2380.

Staub, O. & Rotin, D. (1997). Regulation of ion transport by

protein—protein interaction domains. Current Opinion in Nephrologyand Hypertension 6, 447—454.

Staub, O., Yeger, H., Plant, P. J., Kim, H., Ernst, S. A. & Rotin,

D. (1997). Immunolocalization of the ubiquitin-protein ligase Nedd4

in tissues expressing the epithelial Na¤ channel (ENaC). AmericanJournal of Physiology 41, C1871—1880.

Taglialatela, M., Toro, L. & Stefani, E. (1992). Novel voltage

clamp to record small, fast currents from ion channels expressed in

Xenopus oocytes. Biophysical Journal 61, 78—82.

Tsuchiya, K., Wang, W., Giebisch, G. & Welling, P. A. (1992).

ATP is a coupling modulator of parallel Na,K-ATPase—K-channel

activity in the renal proximal tubule. Proceedings of the NationalAcademy of Sciences of the USA 89, 6418—6422.

Tucker, S. J. & Ashcroft, F. M. (1998). A touching case of channel

regulation — the ATP-sensitive K¤ channel. Current Opinion inNeurobiology 8, 316—320.

Turnheim, K. (1991). Intrinsic regulation of apical sodium entry in

epithelia. Physiological Reviews 71, 429—445.

Wendt-Gallitelli, M. F., Voigt, T. & Isenberg, G. (1993).

Microheterogeneity of subsarcolemmal sodium gradients. Electron

probe microanalysis in guinea-pig ventricular myocytes. Journal ofPhysiology 472, 33—44.

Acknowledgements

We thank Mrs J. Fakan, Mrs F. Voinesco, Miss N. Ruchonnet and

Mr F. Ardizonni from the Centre of Electron Microscopy of the

University of Lausanne for their expert and kind technical help.

We also wish to thank Dr S. Fakan for his helpful comments and

Dr J.-Y. Lapointe for his help in setting up the cut-open oocyte

technique. We are grateful to Drs S. Kellenberger, L. Schild and

O. Staub for sharing unpublished observations and for their critical

reading of the manuscript. This work was supported by the Human

Frontier Science Program, grant RG-0464.

Corresponding author

J.-D. Horisberger: Institute of Pharmacology and Toxicology,

University of Lausanne, Bugnon 27, CH-1005 Lausanne,

Switzerland.

Email: [email protected]

Feedback inhibition of epithelial sodium channelJ. Physiol. 516.1 43