Opposite effects of Ni2+ on Xenopus and rat ENaCs expressed in Xenopus oocytes

Dana Cucu,1 Jeannine Simaels,1 Jan Eggermont,1 Willy Van Driessche,1 and Wolfgang Zeiske2

1Laboratory of Physiology, Department of Molecular Cell Biology, K. U. Leuven, Campus Gasthuisberg O & N, Leuven, Belgium; 2Division of Animal Physiology, Department of Biology/Chemistry, University of Osnabrück, Osnabrück, Germany

Submitted 25 August 2004 ; accepted in final form 30 May 2005


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Opposite effects of Ni2+ on Xenopus and rat ENaCs expressed in Xenopus oocytes. Am J Physiol Cell Physiol 289: C946–C958, 2005. First published June 8, 2005; .—The epithelial Na+ channel (ENaC) is modulated by various extracellular factors, including Na+, organic or inorganic cations, and serine proteases. To identify the effect of the divalent Ni2+ cation on ENaCs, we compared the Na+ permeability and amiloride kinetics of Xenopus ENaCs (xENaCs) and rat ENaCs (rENaCs) heterologously expressed in Xenopus oocytes. We found that the channel cloned from the kidney of the clawed toad Xenopus laevis [wild-type (WT) xENaC] was stimulated by external Ni2+, whereas the divalent cation inhibited the channel cloned from the rat colon (WT rENaC). The kinetics of amiloride binding were determined using noise analysis of blocker-induced fluctuation in current adapted for the transoocyte voltage-clamp method, and Na+ conductance was assessed using the dual electrode voltage-clamp (TEVC) technique. The inhibitory effect of Ni2+ on amiloride binding is not species dependent, because Ni2+ decreased the affinity (mainly reducing the association rate constant) of the blocker in both species in competition with Na+. Importantly, using the TEVC method, we found a prominent difference in channel conductance at hyperpolarizing voltage pulses. In WT xENaCs, the initial ohmic current response was stimulated by Ni2+, whereas the secondary voltage-activated current component remained unaffected. In WT rENaCs, only a voltage-dependent block by Ni2+ was obtained. To further study the origin of the xENaC stimulation by Ni2+, and based on the rationale of the well-known high affinity of Ni2+ for histidine residues, we designed {alpha}-subunit mutants of xENaCs by substituting histidines that were expressed in oocytes, together with WT {beta}- and {gamma}-subunits. Changing His215 to Asp in one putative amiloride-binding domain (WYRFHY) in the extracellular loop between Na+ channel membrane segments M1 and M2 had no influence on the stimulatory effect of Ni2+, and neither did complete deletion of this segment. Next, we mutated His416 flanked by His411 and Cys417, a unique site for possible heavy metal ion chelation, and, with this quality, most proximal (~100 amino acids upstream of the second putative amiloride binding site at the pore entrance), was found localized at M2. Replacing His416 with arginine, aspartate, tyrosine, and alanine clearly affected amiloride binding in all cases, as well as Na+ conductance, as expressed in the xENaC current-voltage relationship, especially with regard to aspartate and tyrosine. However, similarly to those obtained with the WYRFHY stretch, none of these mutations could either abolish the stimulating effect of Ni2+ or reverse it to an inhibitory type.

epithelia; divalent cations; amiloride; Na+; voltage clamp


THE AMILORIDE-SENSITIVE EPITHELIAL NA+ CHANNEL (ENaC) regulates a variety of physiological functions in different Na+-transporting epithelia such as those in the distal nephron, the distal colon, lung epithelia, and duct cells of exocrine glands. The activity of ENaCs is highly regulated by hormones such as aldosterone and vasopressin but also by extracellular factors via mechanisms that are not completely understood. One remarkable mechanism by which the activity of ENaCs is regulated in native Na+-transporting epithelia is called self-inhibition, which can be observed as a peak current that relaxes to a lower value with a time course of a few seconds after a sudden increase in the extracellular Na+ concentration ([Na+]o) (6, 29, 36). Polyvalent cations such as Cd2+, Ni2+, Zn2+, and La3+ were shown to increase Na+ uptake in frog skin (35) by interfering with the self-inhibition process, whereas in oocytes injected with the cRNA of the homologous rodent ENaC subunits, Ni2+ exerted opposite effects compared with those in the native tissues, and some heavy metal ions had no effect at all (25). However, Zn2+ turned out to be a self-inhibition blocker in the mouse ENaC (mENaC) (29). Most intriguingly, in frog skin but also in a cell line derived from the Xenopus kidney (A6), Ni2+ and other divalent cations stimulated active Na+ transport (8, 10), whereas it had inhibitory effects in Xenopus oocytes expressing mENaCs (28) and rat ENaCs (rENaCs) (27).

The use of Ni2+ as an inorganic Na+ channel probe is well suited to the detection of strategic cysteine and histidine residues in the Na+ channel structure. Moreover, the study of toxic effects of this heavy metal, well known to be a very harmful pathogen in technology, will allow for the assessment of risk factors arising from interactions with accessible apical membrane transporters such as the ENaC. In a previous study (8), we used A6 epithelia, an immortalized cell line derived from the distal nephron of the clawed toad Xenopus laevis, to explore the effects of external Ni2+ on Na+ transport. Ni2+ stimulates Na+ transport in A6 epithelia. Amiloride-induced current fluctuation analysis demonstrated competition between Na+ and amiloride on the one hand and between Ni2+ and Na+ as well as amiloride on the other hand. In the present study, to better understand the stimulatory mechanism of Ni2+ on ENaCs and the competition with amiloride and Na+, we investigated the effects of Ni2+ on the channel cloned from Xenopus ENaC (xENaC) A6 cells expressed in Xenopus oocytes.

Moreover, we have investigated whether the effect of Ni2+ is specific for the xENaC or whether stimulation is a general characteristic of ENaCs cloned from other organs and/or species. Therefore, we compared the effect of Ni2+ on rENaCs cloned from rat colon and expressed in Xenopus oocytes. We found that Ni2+ stimulates Na+ current and conductance in xENaCs and has an inhibitory effect in rENaCs (27). The results obtained in rENaCs are also in agreement with previous observations made regarding mENaCs (28). Most interestingly, a recent report by Sheng et al. (29) may have shown that Zn2+, in contrast to Ni2+, was able to stimulate the current through mENaCs and that a cysteine was pinned down as a possible reaction partner. In their former study of Ni2+, these authors identified key extracellular histidine residues ({alpha}-His282 and {gamma}-His239) within conserved regions of the mENaC {alpha}- and {gamma}-subunits that were required for channel block. This segment (WYRFHY) represents a putative binding site for amiloride. To extend our previous experience with rENaCs and xENaCs, we compared Na+ channel characteristics and amiloride binding for the wild-type (WT) channels of both species after expression in Xenopus oocytes. Moreover, in an effort to localize the site (presumably histidine) at which Ni2+ binds in the xENaC to exert its stimulatory effect, we performed mutagenesis studies of the extracellular segment of the {alpha}-subunit of xENaC. We investigated xENaCs (WT and mutated) and WT rENaCs by means of two electrophysiological methods. One method was the classic two-microelectrode voltage-clamp (TEVC) technique. The other approach used was the transoocyte voltage-clamp (TOVC) method recently developed in our laboratory (7). The TOVC method was used for the investigation of transoocyte current (ITO), transoocyte conductance (GTO), and amiloride-induced fluctuation in current. The TOVC method is an excellent technique with which to record current fluctuation spectra, to make accurate estimates of the amiloride association rate constant (kon), and to compare this parameter obtained from the xENaC with that from the rENaC in the WT. Furthermore, we assessed the channel characteristics [i.e., current-voltage (I-V) relationships] as well as the amiloride kinetics before and after Ni2+ treatment and/or histidine mutation.


    MATERIALS AND METHODS
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The research and experiments described the present report were performed with the support of the Belgian Science Foundation (Fonds voor Wetenschappelijk Onderzoek Vlaanderen). The use of Xenopus laevis in the Laboratory of Physiology in our study was approved by the Ministry of Agriculture (approval no. LA1210202).

Preparation of ENaC mRNA and ENaC mutants. Xenopus and rat ENaC cDNA encoding the {alpha}-, {beta}-, and {gamma}-subunits (gift of B. Rossier and J. D. Horisberger, University of Lausanne, Lausanne, Switzerland) were cloned into the pSDEasy and pSport 1 vectors, respectively. Point mutations were generated in {alpha}-xENaC cDNA with the sequential polymerase chain reaction (PCR) method using Pfu DNA polymerase (QuickChange site-directed mutagenesis kit; Stratagene, La Jolla, CA).

Expression of ENaCs in Xenopus oocytes. Xenopus females were purchased from the African Xenopus Facility (Knysna, South Africa). They were anesthetized by inducing hypothermia, and the ovarian lobes containing oocytes were removed. Oocytes were defolliculated by incubation in collagenase (1 mg/ml; Serva, Mannheim, Germany) for 2 h and subsequently washed with Ca2+-free Ringer for 10 min (see below for composition). cRNA of each of the {alpha}-, {beta}-, and {gamma}-subunits were synthesized, and equal amounts of subunit cRNA (5 ng of total cRNA) were injected into oocytes.

Solutions and chemicals. Native noninjected oocytes were incubated in Ringer solution containing (in mM) 90 NaCl, 2 CaCl2, 3 KCl, and 5 HEPES, pH 7.6. For the storage of ENaC-injected oocytes, we used a low-Na+ Ringer solution that contained (in mM) 5 NaCl, 85 N-methyl-D-glucammonium Cl (NMDG-Cl), 2 CaCl2, 3 KCl, and 5 HEPES, pH 7.6. The experiments were performed in solutions with the following composition (in mM): 102 NaCl, 2.5 KHCO3, and 1 CaCl2, pH 8. NiCl2 (2 mM) was added to the solutions without osmolality adjustment.

Transoocyte voltage clamp. Transoocyte current and conductance from ENaC-expressing oocytes were measured using the TOVC technique as described previously (7). Briefly, the oocyte is mounted in a container designed to fit in an Ussing-type chamber. Figure 1A shows that one side of the oocyte was exposed to 102 mM NaCl-Ringer solution; this is referred to as the high-Na+ (HN) side, whereas the other side was exposed to Na+-free solution and is referred to as the zero-Na+ (ZN) side (NMDG-Cl-Ringer, no Na+). In the TOVC arrangement, the positive current indicates cation movement from the HN side to the ZN side.



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Fig. 1. Experimental procedures. A: schematic arrangement of the oocyte chamber with the transoocyte voltage-clamp (TOVC) method. The oocyte was mounted in an electrically well-sealing container in the middle of a modified Ussing chamber (for details, see Ref. 7). Wild-type (WT) or mutated epithelial Na+ channel (ENaCs) from Xenopus distal kidney (A6 cell culture) or rat colon were expressed in the oocyte. One Ussing chamber compartment was perfused with a high-concentration (102 mM) Na+ saline (HN; left), and the other was perfused with a Na+-free or 0 Na+ saline (ZN; right), where NaCl had been replaced by N-methyl-D-glucammonium Cl (NMDG-Cl), thus establishing a HN-to-ZN-directed Na+ concentration ([Na+]) gradient and driving the Na+ flux. The electrical equipment consisted of a four-electrode voltage-clamp arrangement because it is typically used for epithelial recordings to measure and/or impose the transepithelial voltage (VTO) and allow for the measurement of the transoocyte current (ITO), which equals the short-circuit current (Isc) at VTO of 0 mV. The clamp apparatus also enabled fluctuation analysis of Na+-carried currents induced by the ENaC blocker amiloride. B: whole cell Xenopus laevis ENaC (xENaC) current response to a voltage pulse ({Delta}V) obtained with the two-electrode voltage-clamp (TEVC) method, which comprises time-dependent development of instantaneous current (Iinst) and voltage-activated current phases ({Delta}IV). An exponential fit of the amiloride-sensitive current was performed for the data points below the dotted line after the establishment of Iinst. {Delta}IV is the amplitude of the exponential fit. The current-generating voltage pulse was –140 mV, cytosol negative. The fitted curve is superimposed on the data. The initial current jump was Iinst = –7.0 µA and {Delta}IV = –9.8 µA.

 
Transoocyte current and voltage were recorded with a digital signal processor (DSP) board (model 310B; Dalanco Spry, Rochester, NY) equipped with two high-speed analog-to-digital converters (14 bit) and two digital-to-analog converters (12 bit). A second DSP board was used to record the transoocyte resistance (RTO) simultaneously by continuously imposing low-frequency sine wave voltage (1 Hz) to the oocyte. RTO was calculated as the ratio of amplitude of the voltage and current response. Transoocyte conductance was GTO = 1/RTO.

This technique allows measurements of transoocyte current and conductance as well as current fluctuation analysis (7). For current fluctuation analysis, the HN side of the oocyte was exposed to different amiloride concentrations for short periods. Current noise was amplified, digitized, and Fourier-transformed to yield power density spectra. We recorded noise spectra as the mean of 50 sweeps of 2-s duration, resulting in a fundamental frequency of 0.5 Hz. The interaction of amiloride with the Na+ channel induced a Lorentzian component in the power density spectra. The Lorentzian parameters, the low-frequency plateau (So), and the corner frequency (fc) were determined using nonlinear curve fitting of the spectra (see Fig. 2 and Ref. 32). The on (kon) and off (koff) rate constants of the blocking reaction were calculated from the fit of the following relationship:

(1)
where [B] is the blocker concentration (32). However, statistical weight was assigned only to the easily calculable association rate. Numerous previous investigations (summarized in Ref. 32) showed that the fitted amiloride off rate is too close to the origin of the 2{pi}fc-amiloride concentration relationship, so there is no reliability in the values (cf. Fig. 5A) that may exhibit a small shift after treatment with, e.g., Ni2+, which makes determinations of Ki (koff/kon) from noise analysis critical.



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Fig. 2. ENaC inhibition by amiloride blocker-induced noise in TOVC arrangement. A: amiloride block of ITO and transoocyte conductance (GTO) in xENaCs. Transoocyte potential was +40 mV, and ITO and GTO were recorded in the presence of increasing amiloride concentrations in the HN solution as indicated. B: inhibition of relative ENaC conductance (Grelative) by amiloride for oocytes expressing xENaCs at two [Na+] levels in the HN solution (N = 4, n = 12). Grelative is the ratio of the conductance in the presence of amiloride (Gamiloride) to the value recorded in the absence of the blocker (Gcontrol): Grelative = Gamiloride/Gcontrol. Solid lines represent fits of the data using Eq. 2. The Ki values were 185 ± 20 and 458 ± 10 nM for 30 mM ({circ}) and 102 mM ({bullet}) Na+, respectively. C: power density spectra in the presence of 3 µM amiloride ({circ}) and 8 µM amiloride ({bullet}). Solid lines represent the nonlinear fit of the data by a Lorentzian function plus nonlinear 1/f background component (see MATERIALS AND METHODS). The resulting Lorentzian fit parameters for 3 µM amiloride were corner frequency fc = 15.8 Hz and Lorentzian plateau So = 3.17 x 10–21 A2s/cm2, and for 8 µM amiloride, the same parameters were fc = 33.6 Hz and So = 1.42 x 10–21 A2s/cm2. D: relationship between corner frequency and amiloride concentration. Using linear regression analysis (solid lines), we obtained the following mean ± SE values for [Na+]HN of 30 mM: kon = 33.5 ± 0.9 µM–1·s–1 and koff = 8.9 ± 2.0 s–1 (N = 3, n = 6). At [Na+]HN of 102 mM, we obtained kon = 17.4 ± 1.8 µM–1·s–1 and koff = 9.6 ± 5.3 s–1 (N = 3, n = 8).

 


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Fig. 5. Effect of Ni2+ on the chemical binding rate (2{pi}fc) of amiloride to xENaCs and rENaCs. A: linear regression analysis was used to determine kon and koff according to Eq. 1. With xENaCs, in the absence of Ni2+ ({circ}), kon = 17.8 ± 0.4 µM–1·s–1 and koff = 9.1 ± 3.3 s–1. In the presence of 2 mM [Ni2+] ({bullet}), kon = 7.2 ± 0.3 µM–1·s–1 and koff = 11.6 ± 2.7 s–1 (N = 3, n = 6). In rENaCs, in Ni2+-free solutions ({square}), kon = 21.6 ± 1.2 µM–1·s–1 and koff = 36.7 ± 10.8 s–1, whereas in Ni2+-containing solutions ({blacksquare}), kon = 8.7 ± 0.2 µM–1·s–1 and koff = 5.8 ± 2.2 s–1 (N = 2, n = 5). B: decreasing effect on the amiloride-binding rate at 8 µM HN-amiloride at different [Na+]HN concentrations, with and without 2 mM Ni2+ in the HN solutions in xENaC-expressing oocytes (circles) as well as in rENaC-expressing oocytes (squares).

 
Amiloride-sensitive currents in ENaC-expressing oocytes were obtained by subtracting the currents remaining after 100 µM amiloride addition at the end of each experiment. To analyze titration curves for inhibition of macroscopic Na+ conductance (GNa), the ratio GNa/G0 measured in the presence of amiloride (GNa) to that in the absence of the blocker (G0) is described by a Langmuir inhibition isotherm (22, 25):

(2)
where [B] is the concentration of amiloride, Ki is the inhibitory constant of the blocker, and n' is a pseudo-Hill coefficient.

Two-microelectrode voltage-clamp technique. The measurements were performed using the standard TEVC technique (30). The oocytes were placed in a small Plexiglas chamber and continuously superfused with solutions. Microelectrodes were pulled from borosilicate glass capillaries with a thin filament (Clark Electromedical Instruments, Reading, UK) equipped with a Ag-AgCl wire. Both electrodes were filled with 3 M KCl. Using micromanipulators, the oocyte was impaled with the microelectrodes under a low-magnification stereomicroscope. We used a voltage-clamp amplifier manufactured by Warner Instruments (Hamden, CT). All transmembrane ion currents were measured as a deflection from the baseline current. The ground electrodes in the bath were also made of Ag-AgCl wires. The flow of positive charge (i.e., Na+) from the bathing solution to the oocyte cytosol is termed inward current and is conventionally expressed with a negative sign. I-V relationships were studied with voltage pulses ranging from –140 to +40 mV in increments of 20 mV. Between voltage pulses, the oocytes potential was held at 0 mV. The duration of a pulse was 400 ms, and the interval between pulses was 1,500 ms. The ENaC conductance was calculated by performing linear regression analysis of the data points between –140 and –80 mV.

Analysis of voltage-activated currents. When a voltage pulse was applied (see above), the difference between the baseline current (zero voltage) and the current magnitude reached after 5 ms was termed instantaneous current (Iinst), representing a prompt reaction to the pulse (see Fig. 1B). Starting from this point, the subsequent slow voltage activation in the xENaC current was fitted with an exponential function whose amplitude was termed the voltage-activated current ({Delta}IV).

Statistics. Results are expressed as means ± SE. N is the number of animal donors, and n represents the number of experiments (oocytes).


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Bioelectrical properties of ENaC-expressing oocytes and analysis of amiloride-induced current fluctuation (TOVC method). Xenopus oocytes have become the most frequently used expression system for studying the structure, function, and regulation of ENaC. Until now, experiments with oocytes expressing ENaCs have used the classic TEVC method. This method is very reliable for measurements of whole cell currents and conductance and for studies of I-V relationships. However, one major disadvantage in studying ENaC-expressing oocytes using the TEVC method is the potential intracellular Na+ loading during long-term exposure. Loading the oocytes with Na+ is certainly less pronounced when using the TOVC method, in which the oocytes are exposed to high [Na+] on one side only as described in MATERIALS AND METHODS. The Na+ flux from the HN side to the ZN side was further enhanced by the application of a transoocyte potential of +40 mV, positive on the HN side. Under these conditions, ITO for xENaCs was in the range of 2.1–5.2 µA, and GTO ranged from 20 to 200 µS. Amiloride added to the HN side blocked the current and conductance in a dose-dependent manner. A typical example of an experiment in which the inhibitory effect of amiloride on GTO and ITO was studied is shown in Fig. 2A with [Na+]HN of 102 mM. In the presence of an electrochemical gradient across the oocyte, a leak current around the oocyte cannot be excluded. Indeed, 100 µM amiloride in the HN bath did not block ITO completely but reduced ITO from 3.04 ± 0.94 to 0.48 ± 0.01 µA and GTO from 181.0 ± 38.2 to 25.6 ± 1.5 µS (N = 3, n = 5). Under the assumption that the amiloride-insensitive current passes through a paracellular leak, we estimated the leak conductance as ~15%. The amiloride-sensitive components of ITO and GTO are referred to as INa and GNa and were used for further calculations. We decided to quantify the inhibitory effect of different doses of amiloride by studying the effect on GNa, because the results obtained from the inhibition of INa were more scattered. Figure 2B shows the effect of amiloride on mean values of Grelative that represent the ratio of GNa in the presence of amiloride to GNa in control. We evaluated the dependence of the macroscopic Ki on [Na+] with 102 or 30 mM [Na+] on the HN side. Ki was determined by fitting the Langmuir isotherm equation (Eq. 2) to the data. Figure 2B shows that Ki was strongly [Na+] dependent. Ki values were 185 ± 20 and 458 ± 10 nM at 30 and 102 mM [Na+], respectively. This relationship suggests competition between Na+ and amiloride. Similarly, in our previous study (8), we found competition between Na+ and amiloride for xENaCs in A6 epithelia, with Ki = 126 ± 7 and 204 ± 2 nM for 30 and 102 mM apical Na+ ([Na+]ap), respectively.

The amiloride-induced fluctuation in ITO was analyzed in oocytes expressing xENaCs and rENaCs under conditions in which transoocyte currents were partly inhibited with the diuretic on the HN side. Power density spectra could be recorded with amiloride concentrations ranging from 1 to 20 µM. In this concentration range, Lorentzian noise was markedly greater than instrumentation noise. Figure 2C shows power density spectra recorded from xENaCs for two amiloride concentrations. The averaged values of fc were 13.0 ± 1.5 Hz at 3 µM and 30.7 ± 0.7 Hz at 8 µM amiloride concentration (N = 3, n = 10). The association (kon) and dissociation (koff) rate constants were determined by performing linear regression analysis of the 2{pi}fc amiloride concentration data (Fig. 2D and Eq. 1). The strong reduction of kon at higher [Na+] confirms competition between Na+ and amiloride as suggested in Fig. 2B and as shown in Fig. 2D.

Obviously, Na+ entry at the HN side occurs through the ENaC and is amiloride sensitive. On the other hand, Na+ exit from the cell side to the ZN side could occur not only through the ENaC but also via the endogenous Na+-K+-ATPase (20). The effect of amiloride on outward Na+ currents has been studied in frog skin (31), and the amiloride inhibition constant was calculated from noise analysis data as 0.31 µM for outward Na+ currents compared with 0.19 µM for inward Na+ currents. Outward Na+ currents were also measured using the TEVC method in human ENaC (hENaC)-expressing oocytes (6). These currents could be inhibited by large (50 µM) doses of amiloride, but Ki for amiloride was not determined. We tried using amiloride from the ZN side, but we could measure amiloride-sensitive currents at zero potential in only 4 of 20 experiments. Figure 3A shows an experiment in xENaCs, in which, upon the addition of 100 µM amiloride to the ZN side, ITO was blocked by ~50%. The inability of amiloride to block Na+ exit observed in 16 of 20 experiments may have been caused by different factors. A first possibility is that the exit of Na+ occurs only via the Na+-K+-ATPases on the ZN side. To verify this hypothesis, we added 100 µM ouabain to the ZN side. To avoid Na+ loading, the oocytes were incubated in Na+-free solutions on both sides and were exposed for only brief periods to high [Na+]HN (Fig. 3B). The ITO remained unaffected despite treatment with 100 µM ouabain for ~30 min. This result suggests that the exit of Na+ to the ZN side was not via Na+/K+ pumps. Another explanation could be that the exit of Na+ on the ZN membrane is mediated through other native Na+ transporters in the oocyte membrane. A voltage-dependent and amiloride-insensitive Na+ channel activated by long depolarization was found in the Xenopus oocyte membrane (4). Because we expected cell depolarization during exposure to high [Na+] on the HN side, it is likely that in our experimental arrangement, this Na+ channel was opened and mediated the exit of Na+. Of course, other transporters, if Na+ coupled, could mediate the efflux of Na+ at the ZN side as well.



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Fig. 3. Investigation of the Na+ exit pathway at the ZN side of the oocyte. A: effect of amiloride (Ami) added to the ZN vs. the HN side. Current was recorded when [Na+]HN was stepped from 0 to 102 mM. Amiloride (100 µM) was added consecutively to the ZN and HN sides. The clamp voltage was 0 mV. Before Na+ exposure on the HN side, the oocyte was kept in NMDG-Cl-Ringer solution. B: lack of inhibition of ITO and GTO by ouabain (100 µM) added for 30 min to the ZN side.

 
Ni2+ effects on ENaC-expressing oocytes: wild-type xENaCs vs. rENaCs. We first studied the Ni2+ dose-response relationship of the oocyte conductance after expression of xENaCs or rENaCs. It turned out (data not shown) that Ni2+ stimulated xENaCs as shown previously for A6 cells (8) and blocked rENaCs as reported previously (27). xENaC stimulation occurred with Michaelis-Menten kinetics (Hill coefficient of 0.9, n = 12, N = 3) and a half-maximal Ni2+ concentration ([Ni2+]) of 210 ± 30 µM. Unfortunately, simple saturation kinetics with respect to the inhibitory effect on rENaCs were not that obvious when the data were tentatively fitted with the Langmuir equation (see MATERIALS AND METHODS). The apparent Hill coefficient was 0.6 (n = 9, N = 3), and a half-maximal [Ni2+] could be estimated only with low confidence as 1.3 ± 0.5 mM. It is likely that the maximal dose concentration for inhibition is of the order of 10 mM. However, [Ni2+] >5 mM could not be tested, owing to solubility problems with the saline used.

Figure 4 shows the effects of a saturating dose of Ni2+ (2 mM) on current and conductance measured using the TOVC method for xENaCs and for a submaximal concentration of 2 mM [Ni2+] in the case of rENaCs. Here, too, the HN side contained 102 mM Na+, whereas on the ZN side, the oocyte was bathed in 102 mM NMDG-Cl solution. Figure 4, A and B, shows ITO and GTO in response to a voltage step from 0 to +40 mV (referenced to the ZN side) and the effect of amiloride. In xENaC-expressing oocytes, addition of 2 mM [Ni2+] to the HN side solutions augmented the amiloride-sensitive conductance by ~50 ± 1% (n = 6, N = 4). The striking similarity of the degree of stimulation in xENaC-expressing oocytes and A6 cells (8) suggests that Ni2+ acts on the channel itself and not via cellular signaling mechanisms that most likely differ in these cellular systems. Therefore, the interaction of Ni2+ with other native proteins in A6 does not seem to cause the activation of the current.



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Fig. 4. The effect of Ni2+ on currents obtained with TOVC to study xENaCs and rat ENaCs (rENaCs). A: stimulation of transoocyte current and conductance in response to 2 mM Ni2+ concentration ([Ni2+]) in the HN solution in a xENaC-expressing oocyte. Transoocyte potential was +40 mV, HN side positive vs. ZN side. B: same conditions as in A showing inhibition of ITO and GTO by 2 mM [Ni2+] in rENaC-expressing oocyte. In both experiments, 100 µM amiloride added on the HN side almost completely abolished current and conductance.

 
Contrary to the effect on xENaCs, 2 mM [Ni2+] inhibited the current and conductance in rENaCs by 30 ± 2% (N = 3, n = 5). In xENaC- as well as in rENaC-expressing oocytes, the current recorded after application of Ni2+ is completely inhibited by large doses of amiloride applied on the HN side, which demonstrates that in both cases, ITO is carried by Na+ through the ENaC. The rapid response of both xENaCs and rENaCs to Ni2+ may indicate a site of action located on the extracellular side of the ENaC and thus may exclude any interference with intracellular targets. A previous study (28) showed that mENaCs expressed in oocytes were blocked by external Ni2+, just like in rENaCs. Sheng et al. identified extracellular histidine residues ({alpha}-His282 and {gamma}-His239) within conserved regions of the {alpha}- and {gamma}-subunits that were required for channel block by Ni2+. These residues are located in the extracellular loop of mouse {alpha}- and {gamma}-ENaC subunits within a segment that represents a putative binding site for amiloride (14). However, the authors mentioned that the inhibitory channel interaction with Ni2+ is not competitive with amiloride, contrary to our findings regarding stimulatory Ni2+ for xENaCs in A6 cells (8). To clarify this point and to elucidate the opposite effect of Ni2+ in Xenopus vs. mammalian ENaCs, we performed noise analysis studies of amiloride-induced fluctuations in current at different Na+ concentrations.

The amiloride-induced Lorentzian components in the ITO noise spectra were recorded in the presence and in the absence of Ni2+. The addition of Ni2+ to the amiloride-containing solutions in 102 mM [Na+]HN on the HN side caused a marked diminution of kon in both xENaCs and rENaCs. Figure 5A demonstrates the influence of 2 mM [Ni2+] on the amiloride kon for xENaCs and rENaCs. For xENaCs, the control kon was 17.8 ± 0.4 µM–1·s–1 and is comparable to the value previously recorded in A6 cells as 20.2 µM–1·s–1 (8). In rENaC-expressing oocytes, kon was 21.6 ± 1.2 µM–1·s–1 and thus compares well with both the kon value obtained from A6 cells and that obtained from xENaC-expressing oocytes. Although Ni2+ exerts an opposite effect on the conductance of rENaCs vs. xENaCs, we thus found a similar, competition-like effect on the binding of amiloride. A comparable reduction of kon was obtained for both species, amounting to 59.7 ± 10.2% (n = 5, N = 2) and 59.2 ± 19.5% (n = 6, N = 3) for rENaCs and xENaCs, respectively. The reduction of kon in xENaCs as well as in rENaCs suggests direct competition between Ni2+ and amiloride for a common binding site. Alternatively, it is conceivable that a steric modification of the channel by Ni2+ hinders amiloride to reach its site for specific blocking farther away. Because both external Ni2+ (Fig. 5A) and high [Na+] (Fig. 2D) diminished amiloride kon, we hypothesized that the effect of both cations on amiloride binding occurs through an interaction at an analogous, if not identical, site in both channel species, thus suggesting competition between amiloride, Ni2+, and Na+.

This prompted us to examine the effect of 2 mM [Ni2+] at different [Na+]HN on the rate of amiloride binding. Current noise induced by 8 µM amiloride was recorded from xENaC- and rENaC-expressing oocytes in the [Na+] range between 30 and 102 mM. Figure 5B shows the effect of Ni2+ on the [Na+]HN dependence of the amiloride chemical binding rate. In control conditions, with xENaCs as well as with rENaCs, the amiloride reaction rate recorded with 8 µM diuretic decreased when [Na+] was elevated. Conspicuously, in the presence of 2 mM [Ni2+], the corner frequency became independent of the [Na+], an observation in support of the presumed competition between Ni2+ and Na+. Previously, for A6 cells, we suggested a competition of Ni2+ with a Na+ receptor outside the channel mouth, located in the extracellular segment and probably identical to the site of self-inhibition (8). This assumption was based on experiments similar to those shown in Fig. 5B. The resemblance to the results obtained in oocytes expressing xENaCs is clear, whereas the relationship to the inhibitory effect of Ni2+ observed in rENaC-expressing oocytes contrasts with the stimulatory effect in xENaCs, despite the obvious Na+-Ni2+-amiloride competition. In this case, a blockade of the channel pore by Ni2+ in rENaCs is conceivable if it impedes amiloride to bind to its high-affinity site located in the channel pore and thus blocks the entry of Na+. To further explore the process of the interaction of Ni2+ with both xENaCs and rENaCs, we recorded I-V curves using the TEVC method.

Oocytes were clamped at 0 mV, and step changes in the membrane voltage were applied as described in MATERIALS AND METHODS. Generally, the oocytes were incubated in low-[Na+] solution (5 mM) to avoid cell loading with Na+, and they were exposed to high (102 mM) [Na+]o solutions only for brief periods. The time dependence of the current response to the voltage pulses, as well as I-V relationships could be studied. The superimposed traces of the currents recorded during the voltage pulse protocol are shown in Fig. 6. From these records (Fig. 6A), it is clear that the membrane current in the case of xENaCs consists of two components: 1) an instantaneous current jump that reflects the conductance of the membrane at 0 mV and 2) a voltage-activated current that becomes particularly apparent at high, hyperpolarizing voltages. The instantaneous and the activated currents are completely abolished by amiloride (data not shown), demonstrating that both components represent an increase in the current flowing through ENaCs. To quantify the activation process, the amiloride-sensitive currents were fitted with a single exponent (see Fig. 1B). From this analysis, we obtained the instantaneous current (Iinst), which represents the current jump at the beginning of the voltage pulse and the voltage-activated current ({Delta}IV), determined as the amplitude of the exponential function. These activated Na+ currents may be attributed to a slow (in the range of tenths of a second) process that is caused by the effect of voltage on channel gating. A similar activation of the ENaC by hyperpolarizing voltages was previously reported for both ENaCs from A6 cells (24) and of human origin (3).



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Fig. 6. Current responses from xENaCs and rENaCs to TEVC voltage steps. A: oocytes expressing xENaC measured in 102 mM [Na+], compared with control conditions (CTRL), displayed a clear, slow, secondary activation at increasingly hyperpolarizing voltages. B: only the instantaneous currents from the conditions in A are stimulated by 2 mM [Ni2+]. C: whole cell current in oocytes expressing rENaC. Currents in the absence of Ni2+ do not exhibit activation at hyperpolarizing steps. D: in rENaC, 2 mM [Ni2+] caused an inhibition of the instantaneous current and a strong but slower secondary inhibition at hyperpolarizing voltages.

 
Figure 6B shows the effect of Ni2+ on Iinst and on {Delta}IV at different voltages in xENaCs. The addition of 2 mM [Ni2+] stimulated Iinst. Interestingly, {Delta}IV was apparently unaffected by the divalent cation. Like xENaCs, rENaCs exhibited large, inwardly rectifying Na+ currents (Fig. 6C) that were amiloride sensitive but inhibited by Ni2+. The process of current activation as shown with xENaC currents at hyperpolarizing voltages was not observed in oocytes expressing the rat homolog (Fig. 6C). The absence of current activation by voltage indicates that this process is dependent on the ENaC species as described in previous studies with regard to hENaCs (2) and herein regarding xENaCs, whereas it was less evident in rENaCs (2). The observation made by Awayda (2) was confirmed in our present experiments with rENaCs. The reason why the rENaC is not activated by strong hyperpolarization is not clear. It should be related to some differences in structure among ENaC species (xENaCs and rENaCs), resulting in different gating sensitivity to voltage. As shown in Fig. 6D, 2 mM [Ni2+] caused not only the blockade of the instantaneous current response to the voltage pulse in the case of the rENaC but also a further time- and voltage-dependent (and slower) inhibition within ~100 ms.

Next, we analyzed in more detail the effect of Ni2+ on Iinst and {Delta}IV for xENaCs by constructing the I-V relationships in the presence and absence of the divalent cation. Figure 7, A and B, shows that Ni2+ stimulated practically only Iinst in xENaCs. Furthermore, Ni2+ stimulated Iinst independent of voltage, because the ratio of the current in the presence of Ni2+ over control (IinstNi2+/Iinst) is ~2 at any voltage between –140 and –40 mV, which is shown in Fig. 7A. Together, these data are consistent with the idea of Na+ transport stimulation at a site located outside the voltage-sensing channel pore. Membrane Na+ conductance (GNa) was calculated from the slope of the linear regression analysis of Iinst between –140 and –80 mV (see Fig. 7A). After Ni2+ addition, GNa increased from 40.4 ± 2.3 to 83.8 ± 7 µS. These data confirm the results obtained from measurements in A6 cells (8). The current stimulation by Ni2+ is fast (Fig. 4A), which indicates that this voltage-insensitive site for the stimulatory action of Ni2+ must be easily accessible. Moreover, the process of stimulation of Iinst is not related to the secondary voltage activation of the channel (Fig. 7B).



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Fig. 7. Current-voltage (I-V) relationships obtained using the TEVC method in WT xENaC- and rENaC-expressing oocytes, and changes in this relationship after Ni2+ addition. A: stimulation by 2 mM [Ni2+] of the Iinst for xENaC (N = 3, n = 8). The currents were measured in 102 mM NaCl-Ringer solution. Amiloride-sensitive currents were calculated by subtracting the values obtained in solutions with 100 µM amiloride. By performing regression analysis (solid straight lines) for voltages more negative than –70 mV, we determined the conductances GNa = 40.4 ± 2.3 µS in CTRL and 83.8 ± 7 µS in Ni2+ solutions. B: lack of significant stimulating Ni2+ effect on the voltage-activated current (N = 3, n = 8). C: I-V curves of amiloride-sensitive instantaneous currents in rENaC in control conditions and in the presence of 2 mM [Ni2+] (N = 2, n = 5). For voltages more negative than –70 mV (solid lines), we calculated GNa = 104.4 ± 2.8 µS in the absence of Ni2+ and 62 ± 0.4 µS in the presence of Ni2+.

 
For further comparison of the effect of Ni2+ on xENaCs and rENaCs, we analyzed the instantaneous current in rENaCs at the beginning of the voltage pulse. Ni2+ diminished Iinst at any voltage as shown in Fig. 7C. From the slope of the linear regression analysis between –140 and –80 mV, we calculated GNa = 104.4 ± 2.8 µS in the absence of Ni2+ and 62 ± 0.4 µS in its presence.

Histidine mutations in the extracellular loop of the {alpha}-xENaC in the domains WYRFHis215Y and HKSWGHis416C: stimulation by Ni2+ is unaffected, although amiloride and Na+ interact differently with xENaCs. All of the above results demonstrate that Ni2+ exerts opposite effects on Na+ current in xENaC- and rENaC-expressing oocytes, although the channels have strikingly similar biophysical properties. Moreover, we have demonstrated that in xENaCs (but also in rENaCs), amiloride binding is impaired by Ni2+ and interferes with Na+. This result is most likely related to the fact that the binding of the diuretic and the divalent cations take place in areas that are in close proximity to each other, if not in identical locations. Therefore, our attempts to localize the binding site of Ni2+ were based on data available for the binding site of amiloride. A mutagenesis screen of amino acids preceding the second transmembrane segment of {alpha}-, {beta}-, and {gamma}-ENaC identified the residues {alpha}-Ser583, {beta}-Gly525, and {gamma}-Gly537 that, when mutated, reduced 1,000-fold the channel affinity to amiloride (18). It has been shown that all ENaCs cloned to date share this site for amiloride inhibition. Before as well as after the identification of the critical role of the homologous {alpha}-Ser583, {beta}-Gly525, and {gamma}-Gly537 in amiloride binding, other mutations were made in search of other residues that contribute to amiloride block. Thus the short amino acid segment WYRFHY in the mouse and rat {alpha}-ENaC domain was also proposed to participate in amiloride binding (14, 19). Deletion of this region resulted in a loss of amiloride binding to the channel. Clearly, the interaction between amiloride and the ENaC is complex and may involve other, as yet unidentified residues in the extracellular loop of the ENaC family. Within the WYRFHY segment, from {alpha}- and {gamma}-subunits of mENaCs, the mutation {alpha}-His282 to aspartate or double mutations to arginine {alpha}-His282Arg/{gamma}-His239Arg eliminated the Ni2+ block in oocytes expressing mENaCs (28). Finally, extracellular loop folding might bring the WYRFHY segment close to the {alpha}-Ser583-{beta}-Gly525-{gamma}-Gly537 amiloride-binding motif at the pore entrance and thus form a complex drug-binding pocket.

To examine whether Ni2+ would bind to the homologous segment in xENaCs and thus contribute to Ni2+ stimulation, we substituted the corresponding histidine residue {alpha}-His215 within the WYRFHY segment of the Xenopus {alpha}-subunit with aspartate and coexpressed an {alpha}-His215Asp ENaC with WT {beta}- and {gamma}-ENaCs. We did not observe any significant changes regarding Ni2+ action on amiloride-binding rates in xENaCs in two experiments (data not shown). Not unexpectedly, 2 mM Ni2+ stimulated the amiloride-sensitive current and conductance by ~40%. Assuming that histidine is important for Ni2+ binding, this suggests that the site for Ni2+ binding and stimulation of xENaCs does not involve the histidine residue from WYRFHY, which, in contrast, participates in the inhibitory Ni2+ binding in mENaCs. Even more so, deleting the entire WYRFHY stretch led to the same result (data not shown), indicating that neither His215 nor its immediate surroundings are related to the xENaC stimulation by Ni2+.

In our previous report on A6 cells (8), we attempted to chemically characterize the residues involved in xENaC stimulation by Ni2+. We found that p-chloromercuribenzoate (PCMB), a reagent that binds to cysteine but not the histidine-reactive diethyl pyrocarbonate (DEPC) mimicked the stimulatory effect of Ni2+. However, from the chemical point of view, histidine rather than cysteine may be the most preferred partner with which to form complexes with Ni2+, e.g., in enzymes such as urease (34), and it has also become an important Ni2+-complexing structure (the so-called His tag) in preparative biochemistry. In the extracellular loop of each xENaC subunit, there are several histidine residues: nine in the {alpha}-subunit, eight in the {beta}-subunit, and six in the {gamma}-subunit. However, they are not grouped into histidine-rich motifs as described for enzymes that bind Ni2+, although protein folding could bring two or more histidine residues (possibly including also cysteines), which appear distant in the sequence map, into close proximity to form the final tertiary structure.

We demonstrated with experiments in xENaCs and rENaCs that Ni2+ impaired the binding of amiloride. Therefore, a residue that is located closer to the high-affinity site for the blocker at the channel mouth is also likely to coordinate Ni2+ and might, even if linearly (on the amino acid counting scale) somewhat distant but perhaps able to fold into the amiloride site neighborhood, become a docking site for Ni2+-Na+-amiloride simultaneously. This is the reason why we chose to analyze the function of the xENaC {alpha}-His416 residue that is closest to the amiloride-binding site at the beginning of the M2 domain (16). The position of the amino acid segment containing His416 is shown in Fig. 8. The sequence map shows not only another histidine (His411) in the vicinity of His416 but also an intimate neighbor, a cysteine residue (Cys417). These three residues may interact with Ni2+ to construct a high-affinity site for Ni2+ binding as shown for the permease of Escherichia coli, for example (5). As mentioned above, we previously suggested a cysteine as one possible Ni2+-binding site in the A6 ENaC (8). Figure 8 also shows that the corresponding segment in the rENaC contains no histidines, but we note that it still shares the cysteine position with Xenopus.



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Fig. 8. Sequence alignment of xENaC and rENaC stretches in the {alpha}-subunit loops of the last histidine-containing domain in the prepore amiloride-binding areas. The first amino acid in the respective stretches is shown in parentheses. The mutated xENaC amino acid His416 is shown in boldface type.

 
In the following section of the text, only {alpha}-subunit mutations of xENaCs are described, and they were expressed in oocytes together with WT {beta}- and {gamma}-subunits. We generated different point mutations by substitution of the {alpha}-histidine residue His416 with the amino acids alanine (A), arginine (R), tyrosine (Y), and aspartic acid (D). However, these mutations did not abolish the stimulating effect of Ni2+ or reverse it to an inhibitory type. Nevertheless, each mutation resulted in channels with different sensitivities to Ni2+ (concerning only the impact on the amiloride kon rate) and different I-V behavior. As shown with the use of the TOVC method, all substitutions of {alpha}-His416 diminished amiloride kon in control conditions (Ctrl; Table 1) in the absence of Ni2+ compared with the WT channels. Interestingly, kon of all mutations becomes roughly two-thirds that of the WT. Also as in the WT, the addition of 2 mM [Ni2+] further diminished kon, but the percentage of this effect was dependent on the amino acid substituted as shown in Table 1. For instance, in oocytes expressing {alpha}-His416Tyr-{beta}WT{gamma}WT channels, Ni2+ decreased kon by 27%, which is only half the effect caused by the divalent in WT channels. To the contrary, in channels with {alpha}-His416Ala, Ni2+ decreased the amiloride on rate by 77%. This reduction of kon is ~30% more than in WT ENaCs. Nevertheless, the final kon values in the presence of Ni2+ seem not to differ too much from the WT case. These observations indicate that the mutation-independent effect of Ni2+ on the amiloride association rate constant excludes any hypothetical Ni2+ interaction with the His416 residue. Moreover, because all mutations of {alpha}-His416 clearly affected the amiloride binding even before Ni2+ was added (see Table 1), an allosteric rather than direct histidine involvement in the observed effects may appear likely. Therefore, the question whether those just-described mutations would have an impact on control Na+ conductance (absence of blocker and stimulator) is most interesting. We addressed this question using the TEVC technique also on mutated channels before and after addition of 2 mM [Ni2+] (cf. MATERIALS AND METHODS and Fig. 7 legend). Figure 9 shows the voltage dependence of the Na+-specific Iinst in the presence and absence of Ni2+ for WT and channels with substitutions of {alpha}-His416. Oocytes expressing mutated channels exhibited an instantaneous voltage activation of current at hyperpolarizing pulses such as those expressing the WT channel, although the magnitude of the current activation by voltage was sometimes too small for an accurate fit as, for instance, in the case of {alpha}-His416Tyr-{beta}WT{gamma}WT. After addition of Ni2+, Iinst from oocytes expressing mutated channels was always significantly higher. This increase, known from WT channels, again shows that the site at which Ni2+ binds cannot be His416; thus other residues must participate in Ni2+ coordination. In light of the finding that the degree of stimulation is nevertheless dependent on the type of amino acid substituted for His416, we assume that His416 might influence the I-V relationships not directly but rather allosterically. We noticed that in oocytes expressing, for instance, channels with aspartate instead of His416, the control currents were much smaller in the absence of Ni2+ compared with the WT. Nevertheless, Ni2+ caused a remarkable increase in this current (Fig. 9A). In contrast, substitution of histidine with tyrosine resulted in currents even smaller than those observed with aspartate and was correlated with a low but still clear stimulation of Iinst by Ni2+ (Fig. 9C). To better analyze this process, we calculated the ratio of current stimulated by Ni2+ to the current in control conditions (Iinst). For oocytes expressing the WT channels, this ratio was ~2 at any voltage between –140 and –40 mV. Substitution of histidine with aspartate (Fig. 9A) caused a sixfold increase of IinstNi2+/Iinst at all voltages, although starting from a much lower level. One simple explanation for this result may be that there is now a strong electrostatic interaction between the divalent Ni2+ and aspartate (negatively charged). Why the currents in the absence of Ni2+ are also smaller than in WT remains obscure at present but may reflect a similar strong interaction with the monovalent Na+ and therefore lead to a reduced conductance. For the channels containing as substitutes the small neutral alanine (Fig. 9D) but also the long and mobile, positively charged arginine (Fig. 9B) for histidine, the currents in control conditions as well as in Ni2+-containing solutions are comparable to those observed in the WT (Fig. 9A), suggesting no hindrance by these mutations. With respect to the drastically reduced currents with the histidine-to-tyrosine mutation (Fig. 9C), the bulkiness and thus presumably the immobility of tyrosine as opposed to the mobile aspartate results perhaps in a much-reduced channel accessibility to Na+ and may be the cause of these observations.


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Table 1. Rate constants and Ni2+-induced downward shift for amiloride binding in experiments with wild-type and mutated xENaCs

 


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Fig. 9. Activation by Ni2+ of Iinst measured in oocytes expressing {alpha}-xENaC-mutated subunits together with {beta}- and {gamma}-WT-xENaC. The data obtained with WT-{alpha}-, {beta}-, and {gamma}-ENaC are indicated only in A ({circ}, no Ni2+; {bullet}, 2 mM Ni2+). For all mutations, Ni2+-free solution is denoted {triangleup}, and Ni2+-containing salines are indicated by {blacktriangleup}. Data represent means ± SE. A: results from oocytes containing the {alpha}-His416Asp mutation (N = 2, n = 4). B: {alpha}-His416Arg mutation (N = 3, n = 5). C: {alpha}-His416Tyr mutation (N = 2, n = 3). D: {alpha}-His416Ala-mutated channels (N = 4, n = 10).

 
Thus far, we may conclude that our mutation assay data in Table 1 and Fig. 9 do not support a role of His416 as a putative ligand of Ni2+ but might rather point to an allosteric influence on the mechanisms underlying Na+ conductance and amiloride block. This makes sense because the rENaC, in which the corresponding segment (see Fig. 8) is devoid of histidines, exhibits the very same features in the Na+-amiloride-Ni2+ interaction as does the xENaC (see Fig. 3) although Ni2+ blocks the former whereas it stimulates the latter channel type (see Fig. 4). While Sheng et al. (28) allocated the mENaC block by Ni2+ to the WYRFHY segment, we could not find a role of the corresponding xHis215 (see above). Whether in the His411KSWGHis416Cys417 segment (cf. Fig. 8) His411 or Cys417—one of the many extracellular cysteines—plays a role in Ni2+ stimulation of xENaCs remains to be elucidated.

We also analyzed the effect of {alpha}-His416 substitution on the process of voltage-activated currents. As depicted in Figs. 6B and 7B for control (His416), Ni2+ had no consistent effect on {Delta}IV, and this was also the case for all mutations (data not shown). Therefore we shall not deal with this issue further.


    DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 
The ENaC is an important component of the transepithelial Na+ transport route. The ENaC is responsible for Na+ balance and thus controls extracellular fluid volume, arterial blood pressure, and the regulation of airway surface fluid. While the regulation of channel synthesis and surface density by hormones has been well explored, the control by extracellular factors involved in the regulation of channel kinetics is poorly understood. Regulation of ENaC activity is possible, e.g., by extracellular Na+ (through a phenomenon called self-inhibition) but also by several organic or inorganic cations that seem to interfere with self-inhibition. Each of the three channel subunits has a large, cysteine-rich extracellular loop of as yet unknown function. Although a direct interaction with this extracellular domain of ENaC has not yet been demonstrated for the above-mentioned extracellular modulators, the available data strongly suggest that ENaC behaves as a ligand-gated channel similar to several other members of the ENaC-degenerin family (13).

In the present study, we have provided the first direct comparison of the functional properties and the effect of Ni2+ in heterologously expressed xENaCs and rENaCs. We report significant differences between these channels with respect to Ni2+ sensitivity and the process of voltage activation. Ni2+ ions may become important tools for molecular identification of regulatory sites in ENaCs from various cell types and tissues.

Ni2+ exerts opposite effects on Na+ current in WT xENaCs and rENaCs.

We found in this study that Ni2+ stimulated xENaCs. The stimulatory effect of Ni2+ is in agreement with the findings of previous studies that showed that Ni2+ as well as other polyvalent cations, such as La3+, Zn2+ (36), and Cu2+ (9), stimulated Na+ uptake in frog skin. To the contrary, in toad urinary bladder, divalent cations such as Ca2+, Mg2+, or Ba2+ blocked the ENaC from the external side in a voltage-dependent manner with an estimated Ki of ~200 mM (23). Because all of the above results were obtained from measurements of Na+ transport in amphibian tissues, it is clear that the reported effects might be species dependent. In view of the fact that Ni2+ is only one of many cationic agents that stimulate ENaCs, its action may occur at a site that is not very selective. However, studies conducted at our laboratory, have shown that divalent and polyvalent cations exert different influences on Na+ transport in the very same epithelial tissue. For instance, in A6 epithelia, Zn2+ inhibits Na+ transport with a Ki of 45 µM (1), whereas Ni2+ stimulated INa with a Km of 0.5 mM (8) and Mg2+ has no effect when added on the apical side (15). In rENaCs (and also mENaCs), Ni2+ is a blocker whereas Zn2+ turned out to be a stimulator in mENaCs, obviously counteracting self-inhibition (29). Clearly, the action of heavy metal ions such as Ni2+ must occur at selective binding sites. On the other hand, the action of divalent cations may be influenced by other regulatory proteins specific to each of these tissues. To circumvent this problem, we analyzed the influence of Ni2+ on oocytes expressing ENaCs. The Xenopus oocyte expression system enabled us to study the influence of Ni2+ on ENaCs cloned from A6 cells and from rat colon. In this way, we were able to compare the results obtained from xENaC- and rENaC-expressing oocytes with those from epithelial cells.

Comparison of amiloride noise (TOVC) and whole cell Na+ current (TEVC).

Previous studies of xENaCs and mENaCs have produced conflicting results regarding the relationship between extracellular divalent cations and amiloride. In the case of A6 cells, xENaC direct activation by Ni2+ and competition with amiloride was reported (8). Because the effect of Ni2+ on A6 cells was investigated using the noise analysis technique, we aimed to implement the same method to study amiloride-induced current fluctuation in intact oocytes. In early studies, noise analysis was widely used for the estimation of the amiloride rate constants in Na+-transporting epithelia. More recently, studies of oocytes expressing ENaCs revealed limits for the accurate calculation of the amiloride inhibition constant and the analysis of INa block. The reason why kon and koff rate constants of amiloride are not reported for patch-clamp experiments in ENaC-expressing oocytes may be due to the slow gating mode of ENaCs that complicates the kinetic analysis (17). The blocking events by amiloride are indistinguishable from the channel-closed state because they have about the same duration. Only one group of researchers (17) has reported amiloride rate constants calculated from blockade of rENaCs expressed in oocytes. To determine the association rate of the blocker, these authors used ENaCs formed by {alpha}- and {beta}-subunits only, because this channel is almost constantly open. Therefore, blocking events could be readily distinguished from the rare spontaneous channel closing. However, a precise estimate of the amiloride rate constants in {alpha}-, {beta}-, and {gamma}-ENaC-expressing oocytes measured using the patch-clamp technique has not been reported to date.

We implemented the noise analysis method adapted for the TEVC method as reported by Segal et al. (26). The disadvantage of this technique is that the amplifier noise level is elevated and produces a limited bandwidth. With the development of the TOVC technique (7), we are now able to compare the amiloride rate constants obtained from A6 cells with those obtained from oocytes expressing xENaCs. We calculated kon as 17.9 ± 0. 4 µM–1·s–1 and koff as 9.1 ± 3.3 s–1, which is in excellent agreement with the values obtained from A6 cells. Our results obtained with oocytes indicated that Ni2+ diminishes the amiloride Ki by decreasing the on rate to 7.2 ± 0.3 µM–1·s–1 and increasing the off rate constant to 11.6 ± 2.7 s–1. It is clear that the determination of the off rate constant is relatively inaccurate (see MATERIALS AND METHODS), so more weight should be given to kon, and the conclusions derived from this analysis are fully consistent with those from the study of macroscopic Ki behavior; that is, the amiloride on rate is the parameter that is influenced by Ni2+ and/or Na+. One simple way to interpret the data shown in Fig. 5 (amiloride-Na+-Ni2+ interaction) would be that Ni2+ competes with amiloride and Na+ at the very same site on the channel protein and that this is true for xENaCs and rENaCs. However, we would then expect Ni2+ not to stimulate but rather to block xENaCs as observed with rENaCs. Alternatively, at a site outside the channel pore, Ni2+ might exert an allosteric effect on Na+-amiloride binding. This mechanism could then stimulate Na+ entry by affecting the channel in a way that hinders amiloride from reaching its site, i.e., by establishing an "apparent competition" due to mutual exclusion effects on different binding sites for Na+-amiloride and Ni2+, respectively.

Another important finding of an earlier report (27) and of our present study is that Ni2+ inhibits the amiloride-sensitive current in oocytes expressing rENaC. The blocking effect of Ni2+ is in good agreement with the observations of Sheng et al. (28), who reported that Ni2+ inhibited mENaC current, but with a Ki of ~0.5 mM, or ~1 order of magnitude smaller than that in rENaCs, which might suggest molecularly different blocking sites and/or Ni2+ accessibility in the two species. This is supported by the findings of Sheng et al. (28), who identified histidine in the WYRFHY section as a ligand for the inhibitory Ni2+ in mENaCs. Our data regarding xENaC-WYRFHY could exclude His215 as ligand of the stimulatory Ni2+. Contrary to the Sheng et al. report on mENaCs, our findings regarding both xENaCs and rENaCs indicate that Ni2+ diminishes amiloride kon by 59.7 ± 10.2%, again suggesting possibly allosteric competition between the blocker and the divalent cation. To gain insight into why the effect of Ni2+ on INa is different in xENaCs and rENaCs, we studied the influence of Ni2+ on INa in the absence of amiloride at rapid step voltages between –140 and +40 mV.

An interesting finding from our TEVC study is that in the xENaC, which is sensitive to voltage, only Iinst is stimulated by Ni2+ rather than the voltage-activated part, {Delta}IV. The rENaC, however, behaves in a fully ohmic manner and is inhibited by Ni2+ not only in the instantaneous current jump phase but in a secondary, slow Ni2+-dependent inhibition of the current that becomes apparent and is even more effective at more negative voltages (Fig. 6). A voltage-dependent blockade by Ni2+ of rENaC can be the result of two different mechanisms. 1) Ni2+ blocks the rENaC pore by occupying a site located within the permeation pathway, where membrane voltage drops. 2) Ni2+, rather than obstructing the permeation pathway, stabilizes the closed state of the channel through a mechanism that would be independent of the blocking itself but nevertheless would be dependent on voltage. For instance, in studies of high-voltage-activated Ca2+ channels, investigators have proposed the existence of two binding sites for Ni2+: one accounting for the direct blocking and an additional one that stabilizes the Ca2+ channel-closed state (21).

The species difference in the Ni2+ effect, stimulating the xENaC but inhibiting the rENaC, could also be only an apparent difference; for instance, it might be the result of a different time lag of the processes involved. In xENaC, voltage activation is slow and thus visible in the recordings, whereas in rENaC the process could be much faster and might be terminated already during the settling time of the voltage clamp, within the first few milliseconds after the voltage step is initiated. Because of the lack of evidence, such an idea seems far fetched. We therefore propose a working model in which Ni2+ binds in the extracellular, voltage-insensitive domain of the xENaC, thereby changing the channel allosterically in such a way that amiloride and its competitor Na+ are impaired in binding to their common site. Such a site should also be available in the rENaC to allow the observed interaction of Na+, Ni2+, and amiloride (Fig. 5). At present, we cannot tell whether the sites in question in xENaCs and rENaCs share analogous chemical features. Previous studies showed that Ni2+ affects a variety of ion channels, such as voltage-gated Ca2+ channels (11), voltage-gated Na+ channels (33), P2X receptors (33), and glutamate receptors (33). The mechanism of Ni2+ effects on these membrane proteins is not always clear in detail, but both direct blocking and allosteric processes have been suggested (12, 21).

As demonstrated herein for Ni2+-induced xENaC stimulation but inhibition of rENaC and mENaC stimulation (28), other transporters are affected by Ni2+ as well, e.g., the human aquaporin 3 (37). High doses of Ni2+ predispose to asthma, lung fibrosis, lung cancer, and kidney cancer. Taken together, these observations suggest that Ni2+-related diseases may originate from epithelial effects in the first place.

Lessons from mutation experiments.

More recently, Sheng et al. (28) suggested that a histidine residue from the extracellular WYRFHY domain of the {alpha}-subunit and one from the extracellular domain of the {gamma}-subunit form the inhibitory Ni2+-binding site in mENaCs. The residue from the {alpha}-subunit is highly conserved among the species mouse, rat, and Xenopus, and substitution of the homologous histidine residue from mENaCs with aspartic acid abolished the Ni2+ effect on amiloride-sensitive Na+ currents. We identified this histidine residue in the {alpha}-subunit of the xENaC as His215, and we generated the point mutation His215 to aspartate. As in the WT, in these channels, Ni2+ caused stimulation of ITO and diminution of the amiloride kon by ~60% (experiments not shown). This result indicates that the mutation {alpha}-His215Asp is not involved as Ni2+-binding site in xENaCs. Moreover, complete removal of the {alpha}-WYRFHY sequence likewise did not impede stimulation by Ni2+, suggesting a different binding site for this divalent cation. Finally, we again want to shed light on a recent finding regarding A6 cells (8), in which the thiol reagent PCMB, but not the histidine tracer DEPC, stimulated xENaC-like Ni2+ ions. It therefore is not unlikely that Cys417, the neighbor of the mutated His416, may be involved in Ni2+ binding. On the other hand, there are plenty of cysteines (almost diagnostic for ENaCs) that might be possible Ni2+ targets in the extracellular loop. The world of Na+ channels and heavy metals seems much more complicated than expected.


    GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This work was supported by grants from the Fonds voor Wetenschappelijk Onderzoek Vlaanderen (Project FWO-V G.0277.03) and the Alphonse en Jean Forton Foundation.


    ACKNOWLEDGMENTS
 
We thank Dr. J. D. Horisberger and Dr. B. Rossier for making available the cDNA of xENaCs and rENaCs.


    FOOTNOTES
 

Address for reprint requests and other correspondence: W. Van Driessche, Laboratory of Physiology, Department of Molecular Cell Biology, Catholic University of Leuven, Campus Gasthuisberg O & N, Herestraat 49, Box 802, B-3000 Leuven, Belgium (e-mail: willy.vandriessche{at}med.kuleuven.be)

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.


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