Copper and protons directly activate the zinc-activated channel

Abstract

The zinc-activated channel (ZAC) is a cationic ion channel belonging to the superfamily of Cys-loop receptors, which consists of pentameric ligand-gated ion channels. ZAC is the least understood member of this family so in the present study we sought to characterize the properties of this channel further. We demonstrate that not only zinc (Zn²⁺) but also copper (Cu²⁺) and protons (H⁺) are agonists of ZAC, displaying potencies and efficacies in the rank orders of H⁺ > Cu²⁺ > Zn²⁺ and H⁺ > Zn²⁺ > Cu²⁺, respectively. The responses elicited by Zn²⁺, Cu²⁺ and H⁺ through ZAC are all characterized by low degrees of desensitization. In contrast, currents evoked by high concentrations of the three agonists comprise distinctly different activation and decay components, with transitions to and from an open state being significantly faster for H⁺ than for the two metal ions. The permeabilities of ZAC for Na⁺ and K⁺ relative to Cs⁺ are indistinguishable, whereas replacing all of extracellular Na⁺ and K⁺ with the divalent cations Ca²⁺ or Mg²⁺ results in complete elimination of Zn²⁺-activated currents at both negative and positive holding potentials. This indicates that ZAC is non-selectively permeable to monovalent cations, whereas Ca²⁺ and Mg²⁺ inhibit the channel. In conclusion, this is the first report of a Cys-loop receptor being gated by Zn²⁺, Cu²⁺ and H⁺. ZAC could be an important mediator of some of the wide range of physiological functions regulated by or involving Zn²⁺, Cu²⁺ and H⁺.

1. Introduction

The zinc-activated channel (ZAC) is a ligand-gated ion channel (LGIC) belonging to the Cys-loop receptor superfamily which also includes the nicotinic acetylcholine (nACh), 5-hydroxytryptamine type-3 (5-HT₃), γ-aminobutyric acid type A (GABA_A) and glycine receptors [1]. These classical members of this superfamily play important roles in physiology and pathology and thus constitute interesting drug targets in a wide range of disorders [2–5]. ZAC is the least understood member of this family, partly because the gene encoding for ZAC is a pseudogene in mouse and rat genomes, which has complicated explorations into the physiological function of the receptor [1,6].

ZAC displays very little amino acid sequence similarity with other members in the superfamily, and thus it is classified within its own subgroup, the closest matches to ZAC being the human 5-HT3A, 5-HT3B and nACh α7 subunits that exhibit approximately 15% amino acid sequence identity to ZAC [1,6]. Expression studies have detected ZAC mRNA in human fetal whole brain, spinal cord, pancreas, placenta, prostate, thyroid, trachea and stomach [1,6]. Furthermore, the presence of ZAC mRNA has been demonstrated by RT-PCR in adult human hippocampus, striatum, amygdala and thalamus [6]. The presence of ZAC has also been demonstrated in human hippocampal CA3 pyramidal cells and in the polymorphic layer of the dentate gyrus by immunolocalization techniques [6].

Transition metals are known to be essential for proper functioning of various proteins critical for cellular viability and for numerous cellular processes, and thus the intracellular and extracellular concentrations of metals are tightly regulated by metal binding proteins and metal specific transporters [7,8]. Additionally, metal ions can influence the activity of cells by modulating various ion channels, no more so than within the CNS where zinc (Zn²⁺) and other metals are recognized as neuromodulators of N-methyl-d-aspartate, GABA_A, nACh, 5-HT₃ and glycine receptors [9]. Zn²⁺ is essential for normal prenatal and postnatal development and is especially important for fetal brain function [10]. Levels of Zn²⁺ and other metals also appear to be important in neurodegenerative disorders such as Alzheimer’s disease as well as other neurological disorders that result in neuronal loss, including epilepsy and ischemia [11–14]. Within the periphery, metals play important roles in the proper functioning of gastrointestinal, immune and reproductive systems [10]. The pancreas and prostate accumulate and release abundant Zn²⁺ and impairment of Zn²⁺ signaling is associated with diabetes, infertility and cancer [15]. Hence, given the overlapping location of ZAC and the systems affected by imbalances in metal signaling, the receptor may govern important functions for human health.

In our previous study of ZAC, this channel was shown to spontaneously open (open in the absence of agonist) when expressed in human embryonic kidney 293 (HEK293) cells, and application of Zn²⁺ was found to evoke a reversible inward current [1]. However, the small amplitudes of Zn²⁺-activated currents through ZAC in HEK293 cells hampered the investigation of the properties of the channel. In the present study, we have explored further the functional properties of ZAC expressed in COS-7 cells, another mammalian cell line often employed to study recombinant proteins. We report the discovery of two additional agonists for ZAC (Cu²⁺ and H⁺) and present new findings about the biophysical properties of the receptor.

2. Materials and methods

2.1. Cell culture and transfections

COS-7 cells (ATCC CRL-1651) and HEK293 cells (ATCC CRL-1573) were cultured in Dulbecco’s Modified Eagle’s Medium supplemented with 10% calf serum, 100 i.u./ml penicillin and 100 µg/ml streptomycin at 37 °C in a humidified 5% CO₂, 95% air atmosphere. Cells were electroporated (110 V, 20 ms Biorad Gene Pulser Xcell, Hercules, CA) with cDNA encoding for human ZAC along with GFP cDNAs (in pCDM8). Both cDNAs encoding a myctagged and the un-tagged ZAC were used, and since no differences in agonist-evoked current characteristics were observed between cells electroporated with the two constructs, the data were pooled. Cells were used 24–72 h after electroporation.

2.2. Patch-clamp electrophysiology

The whole-cell configuration of the patch-clamp technique was used to record currents from voltage-clamped COS-7 cells. For agonist concentration–response relationship experiments, cells were superfused, at a rate of 2 ml/min, with an extracellular solution containing (in mM): 140 NaCl, 5 KCl, 1.2 MgCl₂, 2.5 CaCl₂, 10 3-(N-morpholino)propanesulfonic acid (MOPS), 11 glucose and adjusted to pH 7.4 with NaOH. MOPS was used in this study as it is a Good’s zwitterionic buffer that does not form complexes with the metals, unlike the typical HEPES buffer [16]. Borosilicate glass patch pipettes (resistance 2–5 MΩ) contained (in mM): 140 KCl, 2 MgCl₂, 0.1 CaCl₂, 1.1 EGTA, 10 HEPES, and adjusted to pH 7.4 with KOH.

In the experiments on H⁺-activated currents, acidic solutions that were rapidly applied to ZAC-expressing cells consisted of extracellular solutions adjusted to a pH of 7.0 with HCl. Solutions with a pH range from 4.5 to 6.5 were made up fresh on the day of experiments from addition of HCl to a solution containing (in mM): 110 NaCl, 5 KCl, 1.2 MgCl₂, 2.5 CaCl₂, 10 Trisodium citrate dihydrate, 11 glucose. In the ion substitution experiments, Zn²⁺-activated currents were recorded in extracellular solution (E1) where KCl was replaced with additional NaCl and both MgCl₂ and CaCl₂ were reduced to lower their influence upon zinc-mediated current amplitude. E1 consisted of (in mM): 145 NaCl, 0.1 MgCl₂, 0.1 CaCl₂, 10 MOPS, 11 glucose, adjusted to pH 7.4 with measured amounts of NaOH. To determine the permeability of K⁺ relative to Cs⁺ E1 was exchanged with E2 where all NaCl in E1 was replaced with KCl (pH adjusted with KOH). Extracellular solution E1 was exchanged with solution E3 or E4 to determine the permeability of Ca²⁺ and Mg²⁺ relative to Cs⁺ respectively. These solutions consisted of (in mM): 100 CaCl₂ (MgCl₂ for E4), 5 histidine, 10 glucose (pH 7.4). Intracellular recording solution (I1) contained (in mM) 140 CsCl, 2 MgCl₂, 0.1 CaCl₂, 1.1 EGTA, 10 HEPES, adjusted to pH 7.4 with CsOH. The free intracellular calcium concentration was estimated to be 10 nM [17].

Experiments examining the inhibitory properties of Ca²⁺ and Mg²⁺ at ZAC were conducted with recording solutions I1 and E1 (as detailed above). The concentration of extracellular Ca²⁺ was then raised from 0.1 mM to between 1 and 75 mM by the addition of CaCl₂. Studies examining the block of ZAC by Mg²⁺ were conducted by raising the MgCl₂ concentration to 30 mM. Solutions containing the raised concentrations of Ca²⁺ and Mg²⁺ in the presence or absence of Zn²⁺ were focally applied to the cell via a SF-77B fast-step perfusion system (Warner Inst., Hamden, CT).

Agonists were applied to GFP-positive cells via the fast-step perfusion system. All experiments were carried out at 32–33 °C using a recording chamber and in-line perfusion heaters (Warner Inst.). Typically, liquid junction potentials were nulled with an open electrode in the recording chamber prior to experiments. For the ion substitution studies, liquid junction potentials were measured with an open-tipped electrode and corrected post hoc. Cells were voltage-clamped at −60 mV unless otherwise noted.

2.3. Data acquisition and analysis

Currents were recorded with an Axopatch 200B amplifier (Molecular Devices, Sunnyvale, CA), filtered at 2 kHz and digitized at 20 kHz with a Digidata 1320A (Molecular Devices) and analyzed using either Clampfit (pClamp, Molecular Devices) or Graphpad Prism v.4 software (Graphpad Software, Inc., San Diego, CA). All data are expressed as the arithmetic mean ± S.E.M., and, unless stated otherwise, statistical comparisons were made using the student t-test with statistical significance set at p < 0.05.

ZAC-mediated currents exhibit an increase in current amplitude over the course of the experiment [1]. To compensate for this run-up phenomenon during concentration–response experiments 1mMZn²⁺ was applied before the subsequent agonist application. The amplitudes of the agonist-evoked currents were normalized to the current elicited by the prior application of 1 mM Zn²⁺. Concentration– response relationships were fitted by nonlinear regression to a sigmoidal dose–response with variable slope equation (Graphpad).

Time constants for activation and decay of ZAC-activated currents evoked by maximal concentrations of Zn²⁺ (1 mM), Cu²⁺ (30 µM), or H⁺ (10 µM, pH 5) were determined by fitting exponential functions to the corresponding component of the current wave form using the Levenberg–Marquardt algorithm with least squares minimization (Clampfit Ver. 9.2; Molecular Devices). A period of time (approximately 10 min) after the establishment of the whole-cell configuration was allowed before the ZAC-activated currents were used for analysis. This allowed for changes in current waveform that occurred during run-up to stabilize. The activation phase of the macroscopic current (between 10% and 90% of peak current amplitude) was fitted by a single exponential function. The rate of current decay upon removal of Zn²⁺, Cu²⁺ or H⁺ was determined by fitting traces to exponential equations from the start of current decay to a point at which 90% of the current had decayed toward baseline. Decay components were best fit to two or more exponentials. To allow for comparisons, weighted time constants were calculated. Time constant values τ_activation and τ_decay for different agonists were compared for significant differences using a one-way ANOVA analysis and Bonferroni posttest in Graphpad InStat3 (San Diego, CA).

The reversal potential for ZAC-activated currents (E_ZAC) was determined by ramping the membrane potential from −60 mV to 40 mV (within 1 s) in the absence of agonist and during the plateau phase of the agonist-evoked current. Currents generated during a ramp in the absence of agonist were subtracted from ramp currents in the presence of agonist. Permeability ratios for monovalent cations relative to Cs⁺ (P_X/P_Cs) were calculated using the Goldman–Hodgkin–Katz voltage equation:

$$E_{\mathrm{Z}\mathrm{n}}=\frac{\mathrm{R}\mathrm{T}}{\mathrm{F}}\text{ln}\frac{(P_X/P_{\mathrm{C}\mathrm{s}}){[{\mathrm{X}}^{+}]}_{\mathrm{o}}}{{[\mathrm{C}{\mathrm{s}}^{+}]}_{\mathrm{i}}}$$

where E_Zn is the equilibrium potential of Zn²⁺-activated current RT/F is 26.3 mV at 32 °C, [Cs⁺]_i and [X⁺]_o are the calculated activities of internal Cs⁺ and external monovalent ions, respectively. Individual current–voltage relationships obtained during the voltage-ramp were plotted, and a linear fit to points either side of current reversal was used to calculate the equilibrium potential. Equilibrium potentials were averaged. The ionic activities were calculated by multiplying the ionic concentration by the ion activity coefficient (γ_ion). The γ_ion values were estimated to be: γ_K 0.75, γ_Cs 0.72, and γ_Na 0.76 [17].

2.4. Drugs and reagents

Tissue culture reagents were purchased from Gibco (Life Technologies Corp., Grand Island, NY, USA), amiloride was purchased from Tocris (Minneapolis, MN), and all other reagents were from Sigma–Aldrich (St. Louis, MO). Metal ion solutions readily hydrolyze to form metal-hydroxy and metal-oxy polymers. These polymers may not always form a visible precipitate but can be biologically inert by preventing the interaction of metals with proteins or cause the metal to bind nonspecifically to a protein through electrostatic interactions rather than to high affinity binding sites [18]. To prevent this hydrolysis, metal stock solutions were made with ligands at a metal:ligand molar ratio of 1:6. The metal:ligand combinations used were; Zn(II)–histidine, Cu(II)–glycine, and Fe(II)–citrate [19]. When tested alone, the ligands histidine, glycine and citrate did not activate ZAC (data not shown).

3. Results

3.1. Expression of ZAC in COS-7 cells

In our previous study, transient expression of ZAC in HEK293 cells was found to yield small Zn²⁺-activated currents [1]. We therefore examined another cell line to see if it was more suited for ZAC studies. We transiently expressed ZAC in the African Green monkey kidney COS-7 cell line, and performed the experiments at 32–34 °C. Expressing ZAC in COS-7 cells gave consistently larger Zn²⁺-activated currents than those evoked through the receptor in HEK293 cells (50–200 pA at the start of the experiment in HEK293 cells compared to 200 pA–3 nA at the start of the experiment in COS-7 cells using similar experimental conditions).

Initially, the properties of Zn²⁺ on ZAC transiently expressed in COS-7 cells were investigated. The amplitudes of Zn²⁺-activated currents, recorded from ZAC-expressing COS-7 cells voltage-clamped at −60 mV, increased with increasing concentrations of Zn²⁺ (Fig. 1A). At a Zn²⁺ concentration of 3 mM, the activated currents displayed marked desensitization in the continued presence of the agonist. Fitting the concentration–response data with a sigmoidal equation provided an EC₅₀ value of 180 µM and a Hill coefficient of 2.2 ± 0.2 for Zn²⁺ (n = 3–5 cells, Table 1). Thus, the properties of ZAC expressed in COS-7 cells were very similar to those previously determined in HEK293 cells [1].

Fig. 1.

Zn²⁺ as a ZAC agonist. (A) Inward currents evoked by rapid applications of Zn²⁺ to ZAC-expressing COS-7 cells. Zn²⁺ was applied (solid bar) for 6 s to cells voltage-clamped at −60 mV Zn²⁺-mediated currents recorded from COS-7 cells transiently transfected with ZAC. (B) Concentration–response relationship for Zn²⁺ at ZAC-expressing COS-7 cells. Data shown as mean ± S.E.M. generated from 3 to 5 cells.

Table 1.

Agonist properties at ZAC expressed in COS-7 cells.

Agonist	EC50 (pEC50 ± S.E.M.)	Hill coefficient	% Max response (of 1 mM Zn2+)	n
Zn2+	180 (3.7 ± 0.02)	2.2 ± 0.2	100.1 ± 1.5	3–5
Cu2+	4.0a (5.4 ± 0.1)	~3.3	51.8 ± 2.9	3–14
H+	2.3 (5.6 ± 0.1)	3.7 ± 2.5	141.6 ± 9.8	4–6

EC₅₀ values are given in µM with pEC₅₀ ± S.E.M. or pH₅₀ ± S.E.M. (for H⁺) values in brackets. n refers to the number of COS-7 cells, from at least two separate transfections, used to construct the concentration–response relationships.

^aThe EC₅₀ for the Cu²⁺ concentration–response curve was determined from the fit to the rising phase of the curve.

3.2. Ionic permeabilities of ZAC

The ZAC protein shares some identity within the cationic subdivision of Cys-loop receptors, the 5-HT₃ and nACh receptors [1]. Comparing the reversal potentials associated with changes in the ionic composition of intracellular and extracellular solutions we have previously shown that ZAC is impermeable to anions [1]. In the present study, we examined which cations permeate the ZAC ion channel using extracellular solutions of different ionic compositions to compare the permeabilities of the physiological relevant cations Na⁺, K⁺, Ca²⁺, and Mg²⁺ relative to intracellular Cs⁺.

The relative permeability ratios, P_X/P_Cs, of ZAC channels were calculated by determining the reversal potentials of 1 mM Zn²⁺-activated currents (E_Zn) from voltage ramps (Fig. 2A). Reversal potentials were determined in extracellular solutions with differing ionic compositions (see Section 2). Using NaCl-based extracellular solution and CsCl-based intracellular solution E_Zn was −11 ± 2 mV corresponding to a P_Na/P_Cs of 0.65 ± 0.05 (n = 8). When extracellular NaCl was completely replaced with KCl, E_Zn was −8 ± 1 mV corresponding to a P_K/P_Cs of 0.72 ± 0.04 (n = 7). Hence, there was no significant difference in the permeability of Na⁺ and K⁺ (t-test, p = 0.29).

Fig. 2.

Cationic permeability of ZAC. (A). Voltage ramps between −60 mV and 30 mV applied before and during agonist application. Non-specific currents recorded during voltage ramps in the absence of Zn²⁺ were subtracted from currents recorded during voltage ramps applied during the steady state inward current evoked by 1 mMZn²⁺ (solid bar). Insert; overlapping currents evoked by voltage ramps in the absence (gray current) and presence of Zn²⁺ (black current). (B) Representative Zn²⁺-mediated currents recorded from a ZAC-expressing COS-7 cell in extracellular solutions of varying ionic compositions. Zn²⁺ (1 mM) was applied 1/60 s (Zn²⁺-mediated currents are concatenated in the illustration). The major cationic species is indicated above each trace (see Section 2 for details). When all extracellular monovalent cations were replaced with either Ca²⁺ or Mg²⁺ no detectable Zn²⁺-mediated inward current was observed. (C) Current–voltage relationships from a representative ZAC expressing COS-7 cell obtained from voltage ramps applied during the application of 1 mM Zn²⁺. The current obtained with a NaCl-based extracellular solution was completely absent when Zn²⁺ was applied with a CaCl₂-based solution. The lack of outward current at positive potentials indicates that Ca²⁺ is inhibiting the outward flux of intracellular Cs⁺.

When all extracellular monovalent cations were replaced with either Ca²⁺ or Mg²⁺ no detectable Zn²⁺-activated inward current was observed. Subtracting the current during application of voltage ramps in the absence of Zn²⁺ to that in the presence of Zn²⁺ resulted in no current, suggesting that neither inward nor outward current flowed due to inhibition of ZAC by Ca²⁺ and Mg²⁺ (Fig. 2C).

3.3. Channel inhibition by Ca²⁺ and Mg²⁺

We further examined the apparent inhibition of ZAC by Ca²⁺ and Mg²⁺ observed in the experiments presented in Fig. 2B and C, and found that ZAC-activated currents evoked by 1mM Zn²⁺ were inhibited by extracellular Ca²⁺ and Mg²⁺. With a fixed concentration of extracellular Mg²⁺ (0.1 mM), increasing the concentration of Ca²⁺ from 0.1 mM to 75 mM caused a concentration-dependent reversible inhibition of the Zn²⁺-activated currents (Fig. 3A and B). The IC₅₀ value for Ca²⁺ to inhibit ZAC was 9.3 mM (pIC₅₀ = 2.03 ± 0.07, n = 6–9 cells). In order to examine any potential voltage-dependence to the inhibition by Ca²⁺, 1mM Zn²⁺-activated currents were measured at −60 mV and 60 mV. Inhibition exerted by 30 mM Ca²⁺ was the same at both holding potentials, 47 ± 4% of control (n = 9) at −60 mV and 50 ± 10% of control (n = 4) at 60mV (p = 0.83, Fig. 3C). The inhibition of ZAC by Mg²⁺ was examined by co-applying Zn²⁺(1 mM) with Mg²⁺ (30 mM) with a fixed concentration of extracellular Ca²⁺ (0.1 mM). In the presence of Mg²⁺ (30 mM), the Zn²⁺-activated currents were inhibited by 63 ± 0.6% of control (n = 3, Fig. 3A and B). At 30 mM, Ca²⁺ and Mg²⁺ produced a similar degree of inhibition (p = 0.067).

Fig. 3.

Inhibition of ZAC by Ca²⁺ and Mg²⁺. (A) Overlaid representative Zn²⁺-evoked current in the absence (black current) or in the presence of 30 mM Ca²⁺ or Mg²⁺ (gray current). (B) Concentration–inhibition relationship of the Ca²⁺ block (closed circles) of ZAC. Block of Zn²⁺-evoked current by 30 mM Mg²⁺ is also shown (open circle). Data points are mean ± S.E.M., n = 3–9 cells. (C) Voltage-independence of Ca²⁺ block of ZAC. Left: Zn²⁺-evoked currents at holding potentials of 60 mV and −60 mV in the absence (black currents) and presence of 30 mM Ca²⁺ (gray currents). Right: bar graph showing mean inhibition at 60 mV (n = 4) and −60 mV (n = 9). (D) At a holding potential of −60 mV, application of 30 mM Ca²⁺ evoked a reversible outward current representing the inhibition of spontaneous current. Bar graph showing the mean outward Ca²⁺ (30 mM)-evoked current as a percent of the preceding Zn²⁺ (1 mM)-evoked current (n = 3 cells).

The ability of Ca²⁺ and Mg²⁺ to inhibit the current conductance through ZAC could be caused by inhibiting Zn²⁺ binding via a surface charge screening effect on the surface of ZAC [20]. We examined the inhibition by Ca²⁺ of the spontaneous ZAC-mediated current in the absence of agonist. At a holding potential of −60 mV, application of Ca²⁺ (30 mM) evoked a reversible outward current, which was caused by the inhibition of the spontaneous inward current, that was 22 ± 3.5% of a preceding 1 mM Zn²⁺-activated current (n = 3, Fig. 3D). Thus, Ca²⁺ inhibits the channel itself rather than inhibiting the binding of Zn²⁺ to the channel.

3.4. Copper is an agonist of ZAC

Next we examined the putative ZAC activating properties of several additional transition metals. Rapid applications of iron (Fe²⁺), cobalt (Co²⁺), nickel (Ni²⁺), cadmium (Cd²⁺), and aluminum (Al³⁺), all applied at a concentration of 1 mM, did not produce any ZAC-mediated currents (Fig. 4). In contrast, 1 mM Cu²⁺ evoked a large inward current in cells transfected with ZAC (Fig. 5), but not in untransfected COS-7 cells (data not shown). Cu²⁺ evoked currents in a concentration dependent manner up to 30 µM, whereas currents evoked by Cu²⁺ concentrations greater than 30 µM were characterized by reduced amplitudes (Fig. 5A). Thus, the resulting concentration–response relationship of Cu²⁺ was biphasic. The activating component exhibited an EC₅₀ value of 4.0 µM, an estimated Hill coefficient of 3.3, and a maximal response of 52 ± 3% of the response evoked by 1 mM Zn²⁺ (n = 3–14 cells, Fig. 5B, Table 1).

Fig. 4.

The inactivity of other metal ions at ZAC. A selection of other transition metals and aluminum are all inactive at ZAC expressed in COS-7 cells. At 1 mM concentrations, iron (Fe²⁺), cobalt (Co²⁺), nickel (Ni²⁺), cadmium (Cd²⁺), and aluminum (Al³⁺) were either inactive or elicited negligible responses at ZAC. Zn²⁺ was used as reference agonist in the recordings.

Fig. 5.

Activation of ZAC by Cu²⁺. (A) Inward currents evoked by rapid applications of Cu²⁺ to ZAC-expressing COS-7 cells. Cu²⁺ was applied (solid bar) for 6 s to cells voltage-clamped at −60 mV and compared to currents evoked by 1 mM Zn²⁺. (B) Cu²⁺ concentration–response relationship of ZAC relative to current evoked by 1 mM Zn²⁺. Data shown as mean ± S.E.M. generated from 3 to 14 cells.

3.5. H⁺ is an agonist of ZAC

The expression of ZAC in the stomach [1] prompted us to test the putative effects of protons on ZAC signaling. Application of low pH solutions to COS-7 cells expressing ZAC evoked a reversible inward current at −60 mV (Fig. 6A). ZAC was activated by H⁺ in a concentration-dependent manner, displaying a pH₅₀ value of 5.6 ± 0.1, a Hill coefficient of 3.7 ± 2.5, and a maximal response of 142 ± 10% of the response evoked by 1 mM Zn²⁺ (n = 4–6 cells, Fig. 6B, Table 1). This pH₅₀ corresponded to a H⁺ EC₅₀ of 2.3 µM. Analogously to the Zn²⁺- and Cu²⁺-activated currents, an increase in current desensitization was observed in the presence of higher H⁺ concentrations (Fig. 6A).

Fig. 6.

Activation of ZAC by H⁺. (A) Inward currents evoked by rapid application of H⁺ to ZAC-expressing COS-7 cells. Acidic extracellular solutions were rapidly applied (solid bar) for 6 s to cells voltage-clamped at −60 mV and compared to currents evoked by 1 mM Zn²⁺ (at pH 7.4). (B) H⁺ concentration (pH)–response relationship of ZAC relative to current evoked by 1 mM Zn²⁺. Data shown as mean ± S.E.M. generated from 4 to 6 cells.

3.6. Activation and decay kinetics of Zn²⁺, Cu²⁺ and H⁺ activated currents through ZAC

During the characterization of the concentration–response relationships of Zn²⁺, Cu²⁺ and H⁺, it became apparent that high concentrations of the three agonists gave rise to currents characterized by different activation and decay components. Comparison of τ values obtained for the activation and decay phases of ZAC-mediated currents revealed significant differences for the three agonists at maximal concentrations (Fig. 7A and B). Currents activated by Zn²⁺ (1 mM) peaked at later time-points, resulting in higher τ_activation values (τ = 585 ± 70 ms, n = 3) compared to 30 µM Cu²⁺-activated currents (τ = 367 ± 85 ms, n = 3, p < 0.05, unpaired t-test), and 10 µM H⁺ (pH 5)-activated currents (τ = 234 ± 52 ms, n = 3, p < 0.01, unpaired t-test). Decay of peak current amplitude to baseline after H⁺ exposure (τ = 198 ± 33 ms, n = 3) was also markedly faster compared to currents activated by Cu²⁺ (τ = 706 ± 54 ms, n = 3, p < 0.01, unpaired t-test) or Zn²⁺ (τ = 697 ± 89 ms, n = 3, p < 0.01, unpaired t-test).

Fig. 7.

H⁺-evoked currents through ZAC are characterized by faster activation and decay components than Zn²⁺-evoked currents. (A) Time constants for activation and decay of 1 mM Zn²⁺, 30 µM Cu²⁺, and 10 µM H⁺ (pH 5)-evoked currents. The activation phase of Zn²⁺-evoked currents was significantly slower than Cu²⁺- and H⁺-evoked currents. Decay of H⁺-evoked currents was significantly faster than Zn²⁺- and Cu²⁺-evoked currents. n = 3,* = p < 0.05, ** = p < 0.01. (B) Example current traces from independent experiments with comparable peak values for Zn²⁺- (black), Cu²⁺- (light gray) and H⁺-evoked currents (dark gray), normalization to peak amplitudes at activation or the start of current decay, overlaid so that begin of application and wash off are in parallel for all three traces. Note the significant decay of H⁺-evoked currents.

The proton-mediated currents recorded in COS-7 cells expressing ZAC are similar to those mediated by the prokaryotic proton-gated Cys-loop receptor ortholog GLIC in that they show little desensitization [21]. This is in marked contrast to the desensitization observed with acid-sensing ion channels (ASICs). HEK293 cells endogenously express ASICs that exhibit large amplitude, fast activating currents that desensitize in the continued presence of protons. Additionally, ASIC-mediated currents are blocked in the presence of extracellular amiloride [22]. When ZAC is expressed in HEK293 cells, a typical ZAC-mediated current is observed with the application of zinc. However, an application of pH 5.5 sometimes evoked a larger, faster activating, desensitizing current that is more typical of a ASIC-mediated current (Fig. 8A). When H⁺ was co-applied with 100 µMamiloride, only ZAC-like, slow activating non-desensitizing currents were observed (Fig. 8B).

Fig. 8.

Whole-cell currents recorded from HEK293 cells expressing recombinant ZAC and endogenous ASICs. (A) Overlaid currents recorded from a HEK293 cell transfected with ZAC. Rapid application of Zn²⁺ or H⁺ (solid bar) produced different current characteristics. Zn²⁺ application produced slowly activating, non-desensitizing currents, whereas application of H⁺ produced ASIC-like faster activating currents with pronounced desensitization. (B) In the presence of amiloride, an ASIC blocker, application of H⁺ produced a typical ZAC-mediated current.

4. Discussion

The present study elaborates on our original discovery of the human Cys-loop receptor ZAC [1]. We demonstrate that Cu²⁺ and H⁺ also are agonists of ZAC, which consequently should be considered a Zn²⁺, Cu²⁺- and pH-sensing receptor.

Consistent with our previous study of ZAC expressed in HEK293 cells [1], the currents elicited by Zn²⁺, Cu²⁺ and H⁺ through the receptor expressed in COS-7 cells were all characterized by a lack of macroscopic desensitization at most concentrations. Only at high agonist concentrations did macroscopic desensitization become apparent, reducing peak current amplitude and resulting in a bi-phasic agonist concentration–response relationship, which is most notable with Cu²⁺ but also observed for Zn²⁺ and H⁺ (Figs. 1, 5 and 6). Cu²⁺ and H⁺ displayed 40- and 72-fold higher potencies than Zn²⁺, respectively. Furthermore, H⁺ exhibited ~1.5-fold higher efficacy compared to Zn²⁺, whereas the maximal response elicited by Cu²⁺ was only half of that evoked by Zn²⁺ (Figs. 1, 5 and 6).

The responses elicited by the three agonists were characterized by distinct activation and decay components. When maximal concentrations of the agonists were used, Zn²⁺-evoked currents activated slower than both Cu²⁺- and H⁺-evoked currents, whereas the decay from peak amplitudes of H⁺-evoked currents was markedly faster than the decay of Zn²⁺- and Cu²⁺-evoked currents. At these agonist concentrations, the decay phase would have components of transitions out of the desensitized state and deactivation. The differences in channel characteristics observed for the agonists would suggest that ZAC transitions to and from an open state faster when activated by H⁺ than the metal ions. However, in-depth kinetic studies using ultra-rapid agonist application systems followed by kinetic modeling [23] will be needed to further elucidate the gating kinetics of the respective agonists.

ZAC is a cation-selective channel [1], and in the present study the rank order of permeability to monovalent cations was found to be Cs⁺ > K⁺ = Na⁺. This characteristic is very similar to the monovalent cation permeability of 5-HT₃A receptors [17]. In contrast to the Ca²⁺ and Mg²⁺ permeability of 5-HT₃A and nACh α7 receptors [24,25], our data indicate that ZAC is not only impermeable to both divalent cations, but that the ions also inhibit the channel. In previous studies several residues in the outer extracellular (20′) and cytoplasmic rings (−4′) of transmembrane domain 2 (TM2) in 5-HT₃A and nACh α7 receptors as well as the Asp¹²⁷ in the extracellular vestibule of 5-HT₃A have been identified as determinants of the divalent cation permeability of these receptors [24,26–28]. An alignment of the amino acid sequences of ZAC, 5-HT3A and α7 subunits predicts the corresponding residues in ZAC to be Gln (TM2 20′: Asp²⁹³ in 5-HT₃A, Glu²⁸¹ in nAChα7), Arg (Asn²⁶⁹ in TM2 −4′ ring in 5-HT₃A, Asp²⁵⁷ in nAChα7) and Trp (Asp¹²⁷ in 5-HT₃A, Asp¹¹⁹ in nAChα7), and we propose that these substantial differences in the physicochemical properties of the residues at these positions of ZAC are likely to contribute to the impermeability of Ca²⁺ and Mg²⁺. If ZAC was simply impermeable to Ca²⁺ and Mg²⁺, then one would expect to observe an outward Cs⁺ current at positive potentials. The absence of this current when extracellular Na⁺ were replaced with either Ca²⁺ or Mg²⁺ suggests that the divalent cations inhibit ZAC.

The voltage-independent inhibition of Zn²⁺-activated currents caused by Ca²⁺ and Mg²⁺ could be due to a change in the affinity of Zn²⁺ by altering surface potential through the charge screening of residues [20]. However, ZAC is a spontaneously active ion channel [1] that can open in the absence of an agonist. Application of Ca²⁺ alone inhibited ZAC-mediated spontaneous current, similar to the effect seen when the antagonist, tubocurarine, is applied in the absence of Zn²⁺ [1]. The ability of Ca²⁺ to inhibit the channel in the absence of agonist suggests that the divalent cations are inhibiting ZAC not by preventing Zn²⁺ (or other agonists) binding but by physically inhibiting the ion channel.

The sizes and the physicochemical properties of the three agonists identified for ZAC differs substantially from the endogenous agonists of other mammalian Cys-loop LGICs (ACh, serotonin, GABA and glycine). The location and molecular composition of the consensus orthosteric site in these ‘classical’ Cys-loop receptors are highly conserved. Typically, transmitter binding to these receptors is mediated by an ‘aromatic box’ formed by five aromatic residues, where the quaternary ammonium or protonated amino group of the ligand forms a cation-π interaction with an aromatic residue [29]. Although these aromatic residues are not completely conserved throughout the Cys-loop receptor family a certain degree of consensus exists. Thus, it is striking that none of these aromatic residues are conserved in ZAC. In view of this and the markedly different chemical nature of Zn²⁺, Cu²⁺ and H⁺ compared to the organic transmitters, it seems unlikely that an analogous site exists in ZAC. In ongoing studies in our labs we are investigating whether the underlying mechanism for proton-gating of ZAC could be similar to that proposed for the prokaryotic proton-gated Cys-loop receptor ortholog GLIC [30,31].

As neighbors in the periodic table of elements, the physicochemical properties of Zn²⁺ and Cu²⁺ are similar, furthermore their biological functions also overlap. Like Zn²⁺, Cu²⁺ constitutes a structural component of many proteins, functions as a key cofactor for the catalytic activity of numerous enzymes, and modulates the activity of several ion channels [9,32,33]. Extracellular Cu²⁺ concentration is regulated by the cellular uptake mediated by the transporter CTR1 and the Cu²⁺ export from cells by the ATPases ATP7A and ATP7B [32–34]. To our knowledge, ZAC is the first receptor ion channel shown to be directly activated by Cu²⁺.

The discoveries of the proton-gated Cys-loop receptor orthologs GLIC and PBO-5/6 in Gloeobacter violaceus and Caenorhabditis elegans, respectively, suggest a conserved role for these receptors in pH-sensing [21,35]. The proton-evoked ZAC signaling in this study takes place between pH 5 and 6. This would suggest that ZAC could be physiologically relevant in tissues characterized by slightly acidic conditions, such as the stomach or in pathological acidic conditions.

Physiological concentrations of Zn²⁺, Cu²⁺ and H⁺ are notoriously difficult to measure because free ions rapidly associate with various proteins, and it is unclear if bound Zn²⁺, Cu²⁺ and H⁺ are available to modulate ion channels. Prostate fluid has some of the highest Zn²⁺ content outside the brain; 9 mM [36]. In the CNS, concentrations of Zn²⁺, Cu²⁺ and H⁺ are in the low micromolar range but concentrations from vesicle release into the synaptic cleft are in the 10–200 µM range and even higher in neurodegenerative diseases [9,33,37,38]. However, alternative mechanisms to vesicular release of zinc are possible [39,40]. The pH of extracellular solutions are tightly controlled but synaptic vesicles have high proton concentrations (pH 5.7) such that elevated vesicle release during high neuronal activity could lower the pH in the synaptic cleft [41].

In light of the discovery of Cu²⁺ and H⁺ as ZAC agonists the hypotheses about putative physiological functions of this receptor clearly have to be revised. The fact that both Zn²⁺ and Cu²⁺ and protons activate ZAC in physiologically relevant concentration ranges highlights the potential importance of the receptor in various native tissues. As a promiscuous cation channel activated not only by Zn²⁺ but also by Cu²⁺ and H⁺, ZAC could be envisioned to mediate effects by all three agonists in regions known to contain these agonists in considerable concentrations such as different regions of the CNS, stomach, prostate, and pancreas.

This present study has demonstrated that ZAC forms an ion channel gated by Zn²⁺, Cu²⁺, and H⁺ and is non-selectively permeable to monovalent cations. The role of ZAC in Zn²⁺, Cu²⁺, and H⁺ signaling requires further investigation and could be part of an unexplored signaling cascade that may provide new therapeutic strategies for a number of pathological conditions.

Abbreviations

ZAC

zinc-activated channel

LGIC

ligand-gated ion channel

nACh

nicotinic acetylcholine

5-HT

5-hydroxytryptamine

GFP

green fluorescent protein

MOPS

10 3-(N-morpholino)propanesulfonic acid

CTR1

high affinity copper uptake protein 1

GLIC

Gloeobacter violaceus pentameric ligand-gated ion channel

ATP7A, ATPase

Cu²⁺ transporting, alpha polypeptide

ATP7B, ATPase

Cu²⁺ transporting, beta polypeptide