Abstract
We have shown previously that the function of neuronal nicotinic acetylcholine receptors can be modulated by zinc. This modulation varies from potentiation to inhibition, depending on receptor subunit composition and zinc concentration, with the α4β2 and α4β4 receptors displaying the most dramatic potentiation. In this study, we used site-directed mutagenesis to identify glutamate 59 and histidine 162 on the rat α4 subunit as potential mediators of zinc potentiation. By modeling the extracellular domain of the receptor pentamer, we locate these residues to two subunit-subunit interfaces that alternate with the two acetylcholine-binding interfaces. Substitution of a cysteine at either position allows additional reduction of zinc potentiation upon treatment with the methanethiosulfonate reagents N-biotinoylaminoethyl methanethiosulfonate (MTSEA-biotin) and [2-(trimethylammonium)ethyl] methanethiosulfonate. Mutagenesis and methanethiosulfonate treatment are most effective at position 162, and the presence of zinc hinders the reaction of MTSEA-biotin with the substituted cysteine at this position, suggesting that α4His162 participates in forming a coordination site for zinc. Mutagenesis and methanethiosulfonate treatment are less effective at position 59, suggesting that whereas α4Glu59 may be near the zinc coordination site, it may not be participating in coordination of the zinc ion. It is noteworthy that the position of α4Glu59 within the neuronal nAChR is identical to that of a residue that lines the benzodiazepine-binding site on GABAA receptors. We suggest that the zinc potentiation sites on neuronal nAChRs are structurally and functionally similar to the benzodiazepine-binding sites on GABAA receptors.
Nicotinic acetylcholine receptors (nAChRs) belong to a family of neurotransmitter receptors that includes GABA-, glycine-, and serotonin-gated ion channels. Affinity labeling and mutagenesis studies suggest that agonist-binding sites on these pentameric receptors are located at interfaces between subunits (Corringer et al., 2000). On muscle nAChRs, agonist binding sites are located at the interfaces between the αγ and αδ subunit pairs, with the α subunits contributing a “principal component” of several amino acid sequence segments (A, B, and C) and the γ/δ subunits contributing a “complementary component” of several segments (D, E, and F) (Corringer et al., 2000). The structure of the acetylcholine binding protein (AChBP) confirms this model (Brejc et al., 2001; Smit et al., 2001; Celie et al., 2004). The AChBP homopentamer binds agonist at all five interfaces, with each AChBP monomer supplying the principal contribution to one binding site and the complementary contribution to another binding site. Some neuronal nAChRs are also homopentamers (α7) and form agonist-binding sites in a similar manner (Corringer et al., 1995). However, many neuronal nAChRs are heteromeric with some subunits making the principal contribution (α2-α4, α6) and others making the complementary contribution (β2, β4) (Corringer et al., 2000). For these receptors, the formation of an agonist-binding site at the interface between two dissimilar subunits, and the rotational symmetry revealed by the AChBP structure, limits the number of agonist binding sites to two. This raises an interesting question. If only two interfaces are involved in binding agonist, what is the function of the other interfaces?
Ionic zinc is found in neurons throughout the brain, with highest concentrations in the cerebral cortex and limbic areas (Frederickson et al., 2000). Zn2+ is packaged in synaptic vesicles and, upon neuronal stimulation, is released in a calcium-dependent manner (Assaf and Chung, 1984; Howell et al., 1984). Estimates of extracellular [Zn2+] during neuronal activity vary from 10∼40 μM (Li et al., 2001) to 100∼300 μM (Assaf and Chung, 1984; Vogt et al., 2000). Zn2+ modulates the function of neuronal nAChRs (Palma et al., 1998; Garcia-Colunga et al., 2001; Hsiao et al., 2001) and other ligand-gated ion channels (Huang, 1997), suggesting that Zn2+ is a modulator of synaptic activity. Indeed, synaptically released Zn2+ modulates synaptic activity in the hippocampus (Vogt et al., 2000; Ueno et al., 2002).
Some neuronal nAChRs, such as α4β2 and α4β4, are potentiated by micromolar Zn2+ and inhibited by millimolar Zn2+ (Hsiao et al., 2001). Others, such as α3β2 (Hsiao et al., 2001) and α7 (Palma et al., 1998), are only inhibited by Zn2+. Results with chimeric receptor subunits, and the sensitivity of potentiation to diethylpyrocarbonate and changes in pH, indicate that extracellular histidines participate in forming the potentiating Zn2+ binding site(s) (Hsiao et al., 2001). In this study, we combined conventional mutagenesis, the substituted cysteine accessibility method (SCAM), and protein modeling to identify residues that are determinants of Zn2+ potentiation. Our results suggest that neuronal nAChRs bind ACh and Zn2+ at alternating subunit-subunit interfaces.
Materials and Methods
Molecular Biology. Mutations were introduced into rat neuronal nAChR subunits using the GeneEditor in vitro site-directed mutagenesis system (Promega, Madison, WI). All mutations were confirmed by sequencing. Capped cRNA encoding wild-type and mutant subunits was generated using mMessage mMachine kits (Ambion, Austin, TX).
Preparation of Oocytes and cRNA Injection. Oocytes were surgically removed from mature Xenopus laevis frogs (Nasco, Fort Atkinson, WI). The care and use of X. laevis frogs in this study were approved by the University of Miami Animal Research Committee and met the guidelines of the National Institutes of Health. Follicle cells were removed by treatment with Collagenase B (Roche Diagnostics, Indianapolis, IN) for 2 h at room temperature. Stage V oocytes were injected with 0.5 to 10 ng of each cRNA (at a molar ratio of 1:1) in 15 to 50 nl of water and incubated at 18°C in Barth's saline (88 mM NaCl, 1 mM KCl, 2.4 mM NaHCO3, 0.3 mM CaNO3, 0.41 mM CaCl2, 0.82 mM MgSO4, 15 mM HEPES, pH 7.6, and 100 μg/ml gentamicin) for 2 to 7 days.
Electrophysiology and Data Analysis. Current responses were measured under two-electrode voltage clamp, at a holding potential of –70 mV, using TEV-200 voltage clamp units (Dagan, Minneapolis, MN). Micropipettes were filled with 3 M KCl and had resistances of 0.3 to 2.0 MΩ. Current responses, filtered (8-pole, Bessel low pass) at 20 Hz (–3 db) and sampled at 100 Hz, were captured, stored, and analyzed using a Digidata 1322A (Molecular Devices, Sunnyvale, CA) in conjunction with either a Macintosh G3 computer running Axograph 4.6 software (Molecular Devices) or a Pentium III PC running pClamp 8 (Molecular Devices). Oocytes were perfused at room temperature (20–25°C), in a chamber constructed from Tygon tubing (inner diameter, 0.125 inches), with perfusion solution (115 mM NaCl, 1.8 mM CaCl2, 2.5 mM KCl, 0.0001 mM atropine, and 10 mM HEPES, pH 7.2). Perfusion was continuous (except during MTSEA-biotin applications) at a rate of ∼4 ml/min. ACh alone and in combination with Zn2+ was applied diluted in perfusion solution. Some experiments were conducted using an OpusXpress 6000A Parallel Oocyte Voltage Clamp system running OpusXpress 1.1 and Clampfit 9.1 software (Molecular Devices). In these experiments, all perfusion and application of ACh and Zn2+ was handled by the OpusXpress system. All Zn2+-containing solutions were freshly prepared from a 1 M stock of Zn(CH3COO)2. In previous work, no difference was seen between the effects of Zn(CH3COO)2-and ZnCl2-containing solutions (Hsiao et al., 2001). X. laevis oocytes express a Ca2+-activated Cl– channel that can contribute to whole-cell current responses when Ca2+ permeable channels, such as neuronal nAChRs, are activated. In previous work (Hsiao et al., 2001), we found that the extent of zinc potentiation did not vary across a range of holding potentials (–90 to –40 mV). Because the Cl– channel current amplitude varies dramatically over this voltage range, our results led us to conclude that the Ca2+-activated Cl– channel does not underlie or affect measurement of zinc potentiation of neuronal nAChRs expressed in X. laevis oocytes. For this reason, we have chosen not to attempt to inhibit the Ca2+-activated Cl– channel.
Wild-type and mutant α4β4 receptors displayed little or no desensitization in response to the low ACh concentrations used here, allowing measurement of Zn2+ potentiation as follows. Control current in response to ACh was determined from a 1-s average beginning 29 s after initiation of agonist application and compared with a 1-s average of baseline current immediately before ACh application. Current levels during Zn2+ coapplication were determined from a 1-s average beginning 29 s after initiation of Zn2+ application and compared with the control current. Potentiation of wild-type α4β4 and all mutants (with one exception, see below) was determined with 1 μM ACh, which lies between the EC1 and EC6 for each receptor. We have previously found that the extent of zinc potentiation of wild-type α4β4 does not vary across this portion of the ACh dose-response curve (Hsiao et al., 2001). For one mutant (α4-2E59C β4C75S), 1 μM ACh was the EC0.2. For this receptor, we used 5 μM ACh (the EC2) to test potentiation.
Wild-type and mutant α4β2 receptors displayed substantial desensitization upon exposure to ACh, requiring measurement of potentiation or inhibition as described previously (Hsiao et al., 2001). In brief, the initial 30-s ACh response in the absence of Zn2+ was fit to a single exponential decay function. This fit was projected over the next 30 s during which both ACh and Zn2+ were coapplied. The degree of modulation was measured by taking a 1-s average 29 s after initiation of Zn2+ application and comparing it with a 1-s average of the projected response to ACh alone during the same time period. Thus, both Zn2+ and control values were taken 59 s after the initiation of the experiment. Potentiation of wild-type α4β2 and all mutants was determined with 10 μM ACh, which lies between the EC14 and EC24 for each receptor. We have previously found that the extent of zinc potentiation of wild-type α4β2 does not vary across this portion of the ACh dose-response curve (Hsiao et al., 2001). A similar extent of potentiation was also seen at the lower ACh concentration of 3 μM (the EC3) (Hsiao et al., 2001).
In the SCAM experiments in Fig. 4, A to C, Zn2+ potentiation was measured before and after a 2-min incubation with 2 mM MTSEA-biotin (Toronto Research Chemicals, Inc., North York, ON, Canada). MTSEA-biotin was diluted from a dimethyl sulfoxide stock solution into perfusion solution immediately before application. The final DMSO concentration of 0.5% had no effect on ACh responses or zinc potentiation (data not shown). After the incubation, oocytes were rinsed for 5 min with perfusion solution before measuring Zn2+ potentiation. The high concentration of MTSEA-biotin and relatively long duration of the incubation were chosen to ensure saturation. Indeed, application of fresh reagent for a further 5-min incubation failed to cause any further loss of potentiation (data not shown). In some experiments, 1 mM MTSET was used.
In the SCAM reaction rate experiments in Fig. 4D, potentiation of the response of α4-2H162C β4C75S to 1 μM ACh by 100 μM Zn2+ was measured using our standard protocol. 1 μM MTSEA-biotin was then applied for 5 s. The oocytes were then rinsed for 5 min and potentiation was measured again. This process was repeated and the cumulative exposure times used to determine the reaction rates in the presence and absence of 100 μM Zn2+. The concentration of MTSEA-biotin (1 μM) that would yield a measurable reaction rate was determined empirically. Reaction rates were determined by fitting to the single exponential decay equation: Y = Ymaxe–kt, where Y is the potentiation at time t (in seconds), Ymax is the initial potentiation, and k is the pseudo–first-order rate constant. The second-order rate constant was obtained by dividing k by the concentration of MTSEA-biotin (Pascual and Karlin, 1998).
Both ACh and Zn2+ dose-response curves were fit according to the equation I = Imax/[1 + (EC50/X)nH] where I represents the current response at a given concentration of ACh or Zn2+, X; Imax is the maximal response; EC50 is the concentration of ACh or Zn2+ yielding a half-maximal response; nH is the Hill coefficient. Zinc inhibition data (see Results) was fit according to the equation: I = Imax/[1 + (X/IC50)nH] where I represents the current response at a given metal concentration, X, Imax is the maximal current, IC50 is the concentrations of metal yielding half-maximal inhibition, and nH is the Hill coefficient. Data presented in Fig. 3B was fit to a more complex equation that included both a potentiating and an inhibitory site: I = Imin + (Imax – Imin){[1/(1 + (EC50/X)nH)] – [1/(1 + (IC50/X)mH)]}, where I represents the current response at a given metal concentration, X, Imin is the minimal current, Imax is the maximal current, EC50 and IC50 are the concentration of metal yielding half-maximal potentiation and inhibition, respectively, and nH and mH are the Hill coefficients for potentiation and inhibition, respectively.
Data analysis was performed using Prism 3 software (GraphPad, San Diego, CA). Statistical significance was assessed using a two-tailed unpaired t test, or a one-way analysis of variance followed by the Dunnett's post-test, as appropriate.
Molecular Modeling. Sequence alignment of the amino-terminal extracellular region of the rat α4 and β4 neuronal nAChR subunits with the AChBP was performed using the Alignx module of Vector NTI 5 (InforMax, Inc., North Bethesda, MD). The alignments between the AChBP and α4 and β4 had identity values of 20.4 and 19.4%, respectively. Although the sequence identity between the AChBP monomer and amino-terminal extracellular domains of various nAChR subunits is relatively low, the presence of highly conserved ACh binding residues in the AChBP (Brejc et al., 2001), and the nicotinic pharmacology of the AChBP (Smit et al., 2001) suggests that homology modeling of neuronal nAChR extracellular domains using the AChBP structure is appropriate (Le Novere et al., 2002).
Three-dimensional models were constructed using the program Modeller 6 (Sali and Blundell, 1993) on a Silicon Graphics Indigo2 Extreme workstation. The script “model” was used with neuronal nAChR subunit/AChBP alignments. Disulfide bonds in the AChBP template structure were explicitly included during homology model refinement. The amino-terminal extracellular domain sequences of the α4 and β4 subunits were modeled using the AChBP pentamer structure (Brejc et al., 2001) (Protein Data Bank code 1I9B) to get initial coordinates for an α4β4 pentamer (subunit ordering of αβαββ). Ten models were produced with energy refinement handled within the program. Conditions were optimized such that resulting structures exhibited energies in line with current published nicotinic receptor homology models (Le Novere et al., 2002; Everhart et al., 2003). The lowest energy structure was then inspected visually and with Procheck (Laskowski et al., 1993) for inappropriate stereochemistry (clashing side chains, disallowed torsion angles, etc.). In only one case did a residue require manual adjustment using the O software package (Jones et al., 1991). Further minimization was then carried out using the CNS software package (Brünger, 1998), with 20 cycles of conjugate gradient. The CNS minimized structure was then reanalyzed with Procheck to ensure stereochemical soundness. The images in Figs. 1B and 6 were produced using Ribbons (Carson, 1997). Coordinates for the final α4β4 model may be obtained at http://chroma.med.miami.edu/pharm/faculty_Luetje.html.
Results
The effect of 100 μM Zn2+ on current responses induced by low concentrations of ACh varies from inhibition of α3β2 receptors to greater than 5-fold potentiation of α4β4 receptors, with the α3β4 and α4β2 receptors displaying intermediate levels of potentiation (Fig. 1A). The large difference in the effect of Zn2+ on α4β4 and α3β2 receptors suggested a strategy for identifying the site at which Zn2+ binds and potentiates neuronal nAChRs (i.e., mutagenesis of residues present in α4 and β4, but not in α3 and β2).
Amino acid residues that most commonly form Zn2+-binding sites include histidines, free cysteines, aspartates, and glutamates (Glusker, 1991). Our previous work indicated the involvement of extracellular histidine residues (Hsiao et al., 2001), so we used site-directed mutagenesis to test the role of the seven histidine residues unique to the extracellular portions of α4 and β4. We also examined the two unique extracellular free cysteine residues. Our screen included five residues within the amino-terminal extracellular domain of α4 (His2, His61, His104, His109, His162), two residues within this region of β4 (Cys75, His157), one residue within the carboxyl-terminal extracellular portion of α4 (Cys594), and one residue within the carboxyl-terminal extracellular portion of β4 (His469). At some positions, the residue was changed to the residue present at that position in α3 or β2, as appropriate. For α4His2, α4His104, and β4Cys75, the residues present in α3 and β2 might also be Zn2+-coordinating (glutamate, aspartate, and aspartate, respectively). Thus, in addition to testing α4H2E and α4H104D mutants, we also tested the alanine mutants α4H2A and α4H104A. For β4Cys75, we tested a serine mutant. To test the role of α4Cys594, we used a naturally occurring splice variant (α4-2) that lacks this C-terminal cysteine (Goldman et al., 1987). The positions of these residues within an alignment of α3, α4, β2, and β4 are shown in Fig. 1C. To visualize the positions of these residues within the receptor structure, we generated a homology model of the extracellular domain of the (α4)2(β4)3 pentamer using the AChBP (Brejc et al., 2001) as a template structure (see Materials and Methods). Many of these residues are located at the non–ACh-binding subunit-subunit interfaces within the pentamer (Fig. 1B).
We used the α4β4 receptor as our primary target to allow analysis of α4 and β4 residues in the same receptor. The α4β4 receptor is also convenient because it displays the greatest degree of zinc potentiation, expresses well in oocytes, and shows little or no desensitization in response to low ACh concentrations. The role of critical residues in α4 was also confirmed with α4β2 (see below), a major nAChR subtype in the mammalian central nervous system (Corringer et al., 2000). Mutant α4 and β4 subunits were coexpressed in X. laevis oocytes with wild-type β4 or α4 subunits, respectively. ACh dose-response curves, constructed for each mutant receptor, showed that the mutations had little or no effect on ACh sensitivity (Table 1). We used 100 μM Zn2+ to test the effect of each mutation because 100 μM Zn2+ lies at the peak of the biphasic potentiation-inhibition curve for α4β4 (Hsiao et al., 2001). Thus, reductions in maximal potentiation or shifts in the curve would be detected. Compared with wild-type α4β4 receptors, some mutant receptors (such as α4H162Gβ4) displayed reduced potentiation by 100 μM Zn2+ (45 ± 4% of wild-type), whereas others (such as α4β4C75S) displayed potentiation similar to that of wild-type α4β4 (Fig. 2A). In addition to α4H162G, the α4H61N, and β4H469Y mutations also caused significant decreases in potentiation by 100 μM Zn2+ (Fig. 2B, Table 2). The α4-2β4 receptor (which lacks the C-terminal cysteine) displayed an increase in potentiation, suggesting potential involvement in the allosteric pathway. However, because we are focused on identifying the zinc potentiation site, we chose not to pursue this possibility. In our model of the α4β4 extracellular domain, the side chains of β4Asp195 and α4Glu59 are near the side chain of α4His162 (Fig. 1B). Because Asp195 is conserved in both the β2 and β4 subunits, we substituted an alanine at this position. We also prepared a α4E59A mutant. The α4E59Aβ4 receptor displayed significantly reduced potentiation by 100 μMZn2+, whereas the α4β4D195A receptor did not differ from wild-type α4β4 (Fig. 2B). The results of this mutagenesis screen identify α4Glu59, α4His61, α4His162, and β4His469 as candidates for involvement in forming the zinc potentiation site.
In Fig. 3A, we examined the effects of a range of zinc concentrations on wild-type and mutant α4β4 receptors. The α4E59A, α4H61N, α4H162G, and β4H469Y mutations each reduced maximal potentiation by Zn2+, but none of the mutations increased the EC50 for zinc potentiation, as might be expected if the mutations were damaging a zinc-binding site (Table 2). However, a simple rightward shift in the zinc dose-response curve could be expected only if zinc potentiation were occurring in isolation. We have previously shown that zinc both potentiates and inhibits these receptors at separate sites. At and above 300 μM Zn2+, receptor inhibition of the α4β4 receptor becomes apparent, leading to a distinctly biphasic Zn2+ dose-response curve (Hsiao et al., 2001). If the mutations were causing damage to the potentiation site but leaving the inhibition site intact, a rightward shift in the potentiation curve could be obscured.
To investigate this issue in more detail, we examined the effects of a wider range of Zn2+ concentrations on the α4H162Gβ4 mutant (Fig. 3B). Obtaining accurate values from fitting biphasic zinc dose-response data to a two-site equation is difficult because of the close proximity of potentiating and inhibiting phases (Hsiao et al., 2001). However, in this earlier study, we found that diethylpyrocarbonate treatment eliminates potentiation, without affecting inhibition, of the α4β4 receptor. We fit these data to a single-site inhibition equation to obtain an IC50 for Zn2+ inhibition of 362 ± 70 μM (see Materials and Methods). We then fit the biphasic zinc dose-response data for wild-type α4β4 in our earlier study to a two-site equation (see Materials and Methods) using this IC50 value as a constant. This allowed us to estimate parameters for zinc potentiation of the wild type α4β4 receptor (EC50 = 110 ± 33 μM, nH = 0.96 ± 0.04, maximal potentiation = 1400 ± 250%). The EC50 value is 4-fold greater and the maximal potentiation value 2-fold greater than what we obtain when we fit the potentiating phase of the data to a single site equation. This suggests that the inhibiting phase does indeed partially obscure the potentiation phase. Because zinc inhibition occurs at a separate class of site (Hsiao et al., 2001), it is unlikely to be affected by mutations at the zinc potentiation site. This allows us to use the inhibition parameters obtained from fitting the postdiethylpyrocarbonate treatment wild-type α4β4 receptor when fitting the data obtained for the α4H162Gβ4 mutant. If we also assume that maximal potentiation is not changing, we find that upon fitting to a two-site equation, the EC50 for zinc potentiation of the α4H162Gβ4 mutant is greater (267 ± 7 μM) than the value for wild-type α4β4. This analysis suggests, but does not prove, that the α4H162Gβ4 receptor is indeed less sensitive to zinc potentiation than wild-type α4β4. To prove this conclusively using dose-response analysis, we would need to be able to examine potentiation in the absence of inhibition. However, we have been unable to eliminate, or even damage, the inhibition of nAChRs by zinc (Hsiao et al., 2001). None of the mutations we have examined reduces inhibition by high concentrations of zinc (data not shown). Thus, we turned to other approaches to provide additional information about the role of candidate residues in forming the zinc potentiation site.
First, we examined the effect of double mutant combinations, reasoning that simultaneous mutation of two residues involved in mediating zinc potentiation should yield an effect greater than with either single mutation. Because the α4H162G mutant displayed the greatest loss of Zn2+ potentiation, we used this receptor as a point of reference. Each of the other three mutations was examined as a double mutant with α4H162G. The α4H61N and β4H469Y mutations failed to alter potentiation of the α4H162G β4 receptor. Potentiation of these double mutants by 100 μM Zn2+ was 107 ± 7 and 99 ± 7% of the potentiation of the α4H162G single mutant, respectively. This result suggests that these residues might not play important roles in mediating Zn2+ potentiation. In contrast, receptors formed by the double mutant α4E59A,H162G displayed significantly decreased potentiation compared with the potentiation of α4H162G (81 ± 5% of the single mutant potentiation, p < 0.05, n = 13). It is noteworthy that potentiation was not completely eliminated. Receptors formed by the α4E59A,H162G double mutant retained a modest ability to be potentiated by Zn2+ (36% of wild-type; see Discussion).
As an alternative approach to testing the role of candidate residues in forming a zinc potentiation site, we turned to SCAM (Karlin and Akabas, 1998). In SCAM analysis, a cysteine is placed at the position of interest and function is then measured before and after reaction with a methanethiosulfonate (MTS) reagent. Although free cysteines can participate in coordination of Zn2+, they often do not substitute effectively for other coordinating residues (Paoletti et al., 2000), probably because of the stringent spatial requirements for the relevant atoms (S, N, and O) to be able to coordinate the zinc ion. The result of our double-mutant experiments (see above) suggest that even if a substituted cysteine failed to directly participate in Zn2+ coordination (thus constituting a single mutant), the functional damage to the site would be partial. A further reduction of Zn2+ potentiation after reaction with an MTS reagent would then suggest a physical proximity to the zinc ion and would strengthen the case for direct coordination. As the MTS reagent for most of our experiments, we used MTSEA-biotin, a large, relatively membrane impermeant MTS reagent that has been used to characterize various sites on GABAA receptors (Teissere and Czajkowski, 2001; Wagner and Czajkowski, 2001). To avoid the potential for confounding effects of the MTS reagent acting at other free cysteines, the α4-2 β4C75S mutant receptor, which lacks extracellular free cysteine residues, was used as a “pseudo–wild-type” background (Karlin and Akabas, 1998) in which to test each cysteine mutant. This pseudo–wild-type receptor displayed ACh sensitivity and Zn2+ potentiation similar to that of true wild-type α4β4 and was unaffected by MTSEA-biotin treatment (Tables 1 and 2; Fig. 4, A and C).
We prepared E59C, H61C, and H162C mutants, each within the context of the pseudo–wild-type receptor. Each mutation resulted in significantly reduced Zn2+ potentiation (61 ± 5, 63 ± 9, and 77 ± 2% of pseudo–wild-type potentiation, respectively), similar to what was seen with the E59A, H61N, and H162G mutations. MTSEA-biotin treatment (2 mM, 2 min) had no significant effect on the ACh responses of any of the mutant receptors (data not shown). MTSEA-biotin also had no effect on Zn2+ potentiation of the α4H61C receptor (Fig. 4C), again suggesting that α4His61 may not play an important role in mediating Zn2+ potentiation. In contrast, MTSEA-biotin treatment significantly reduced potentiation of the α4E59C and α4H162C receptors, suggesting that α4Glu59 and α4His162 are located at or near the Zn2+-binding site (Fig. 4, B and C). However, as was the case for the double mutant, these MTSEA-biotin treated mutant receptors retained some ability to be potentiated by Zn2+. In Fig. 4, we used 1 μM ACh to test potentiation of the pseudo–wild-type receptor, as well as the α4H61C and α4H162C receptors. However, we used the equipotent concentration of 5 μM ACh to test potentiation of the α4E59C (see Materials and Methods). We also tested the ability of MTSEA-biotin to reduce zinc potentiation of the α4E59C using 1 μM ACh, finding post-treatment potentiation to be 76 ± 4% of pretreatment potentiation (p < 0.05, n = 6). Zinc potentiation could also be reduced by MTSET application (1 mM, 2 min). Although the trimethylammonium ethyl group deposited by MTSET is substantially smaller than the biotinylaminoethyl group deposited by MTSEA-biotin, it does have a positive charge that would be likely to interfere with Zn2+ binding. Post-treatment potentiation of α4-2H162C β4C75S by 100 μM Zn2+ was 71 ± 3% of pretreatment potentiation (n = 6, p < 0.001) and for α4-2E59C β4C75S, the value was 83 ± 3% of pretreatment potentiation (n = 3, p < 0.05). The effects of MTSEA-biotin and MTSET on zinc potentiation of the α4E59C mutant were substantially less than the effects on the α4H162C mutant. This may be because substitution of a cysteine for α4Glu59 had a greater effect on zinc potentiation than did substitution of α4His162 (61 ± 5 and 77 ± 2% remaining potentiation, respectively). Thus, there is less remaining potentiation to be affected by the MTS reagent. However, it is also possible that although α4Glu59 may be near the site of zinc binding, it might not be directly participating in coordination of the zinc ion.
SCAM can also be used to obtain information about the relative accessibility of a site under different conditions (Karlin and Akabas, 1998). By alternating short exposures to low concentrations of MTS reagent with functional measurements, a reaction rate (and thus a relative measure of accessibility) can be determined (Pascual and Karlin, 1998). In Fig. 4D, we use this method to examine the relative accessibility for MTSEA-biotin at position 162 of α4 in the presence and absence of Zn2+. Oocytes expressing α4-2H162C β4C75S were exposed to 1 μM MTSEA-biotin in 5-s increments. After each application, the oocytes were rinsed, and the extent of potentiation by 100 μM Zn2+ was determined. The decline in Zn2+ potentiation upon repeated exposure to MTSEA-biotin was then fit to an exponential decay function. In the absence of Zn2+, the half-time of the reaction was 8.7 s, yielding a rate of 79,000 ± 14,000 M–1s–1. In the presence of 100 μM Zn2+, the halftime of the reaction was 21.3 s, yielding a rate of 32,000 ± 10,000 M–1s–1. Despite the change in reaction rate, the extent of the effect was the same in the presence and absence of Zn2+ (the fit plateau was 0.69 ± 0.05 and 0.65 ± 0.02, respectively). The significant decrease in reaction rate (p < 0.01) indicates that the site is less accessible to MTSEA-biotin when Zn2+ is present and suggests that Zn2+ is competing with MTSEA-biotin for occupation of the site.
In addition to potentiation of the ACh response, zinc potentiates the response of neuronal nAChRs to nicotine. Potentiation of α4β4 by coapplication of 100 μM Zn2+ with 1 μM nicotine was 444 ± 43% of the response to nicotine alone. The α4H162G and α4E59A mutant receptors each displayed significantly reduced potentiation by 100 μM Zn2+ (50 ± 5% of wild-type α4β4, p < 0.05, and 88 ± 4% of wild-type α4β4, p < 0.01, respectively).
We also examined the effect of mutating α4His162 and α4Glu59 in a different receptor subunit context: α4β2 (Fig. 5). The α4β2 is a major nAChR subtype in the central nervous system (Corringer et al., 2000). The effect of zinc on α4β2 receptors was substantial, with a maximum potentiation of approximately 2.5-fold achieved at 50 μM Zn2+ (Figs. 1A and 5A, and see Hsiao et al., 2001). The α4E59A and α4H162G mutations were each able to significantly reduce the extent of potentiation by 50 μM Zn2+ (Fig. 5B).
Discussion
We have identified α4Glu59 and α4His162 as determinants of Zn2+ potentiation on rat neuronal nAChRs. The positions of these residues within our receptor model are shown in Fig. 6. A strong case can be made for direct coordination of the zinc ion by α4His162. Mutation of this residue results in a large loss of zinc potentiation; MTSEA-biotin has a substantial effect when a cysteine is at this position, and zinc can slow the reaction of MTSEA-biotin with a cysteine at this position. Although the side chain on α4Glu59 is close enough to the side chain of α4His162 (in Fig. 1B, the ϵN of His162 is 5.6 Å from the nearest carboxyl oxygen in Glu59) for a role in coordination to be plausible (Harding, 2001), the effects of mutation and MTSEA-biotin treatment are substantially less than what is seen at position 162. Thus, although α4His162 is likely to be participating in direct coordination of the zinc ion, α4Glu59 may only be near the zinc potentiation site without actually participating in zinc coordination.
The Location of Zinc Potentiation Sites on Neuronal nAChRs. The positions of α4His162 and α4Glu59 within the receptor and the involvement of both α and β subunits in mediating Zn2+ potentiation (Hsiao et al., 2001) suggest that neuronal nAChRs bind Zn2+ at subunit-subunit interfaces that alternate with ACh-binding interfaces. This depends on our decision to model the receptor with a subunit arrangement of αβαββ. Whether the fifth subunit is β or α is irrelevant. In either case, the receptor would have two ACh-binding and two zinc-binding interfaces. More important is the arrangement of the first four subunits in an alternating αβαβ pattern. This arrangement seems to be required. The subunits of the AChBP are arranged in a rotationally symmetric manner, with each subunit supplying a principal face to one ACh binding site and a complementary face to another ACh binding site (Brejc et al., 2001; Celie et al., 2004). In heteromeric neuronal nAChRs, the principal residues are only present on the α subunit, whereas the complementary residues are only present on the β subunit (Corringer et al., 2000). Thus, both α and β subunits are required to form an ACh binding site. Regardless of whether the α/β stoichiometry is 2:3 or 3:2, only two nonadjacent interfaces will form ACh binding sites. An additional two nonadjacent interfaces would form Zn2+-binding sites.
Neuronal nAChRs also form as homopentamers of the α7 subunit (Corringer et al., 2000). The α7 subunit lacks both His162 and Glu59 (Le Novere and Changeux, 2001), and α7 homopentamers are thought to bind ACh at all five interfaces (Corringer et al., 2000). As would be expected, α7 homomers are not potentiated by Zn2+ (Palma et al., 1998). A novel Zn2+-activated channel (ZAC) has been reported (Davies et al., 2003). This homomeric receptor is structurally homologous to nAChRs. Whereas ZAC subunits have a histidine in a position similar to that of α4His162, no aspartates or glutamates are located near the position analogous to α4Glu59. It is unclear whether the ZAC binds Zn2+ at a site similar to the potentiation site we have identified on neuronal nAChRs.
We have identified at least one residue (α4His162) as part of a pair of identical Zn2+ potentiation sites on neuronal nAChRs. Because Zn2+-binding sites can have four, five, or six coordination points (Glusker, 1991), additional residues remain to be identified. These receptors may also possess a second class of Zn2+ potentiation site. In our single-mutation experiments, only partial losses of potentiation were observed. This was not unexpected. However, even the double mutant α4E59A,H162G retained a modest ability to be potentiated by Zn2+ application (approximately one third of wild-type potentiation). In our SCAM experiments, we also did not completely eliminate potentiation. This contrasts with the complete elimination of potentiation upon diethypyrocarbonate treatment (Hsiao et al., 2001). Thus, neuronal nAChRs might possess at least three distinct classes of Zn2+ binding site: a major potentiation site accounting for roughly two thirds of the potentiation (identified here), a minor potentiation site accounting for roughly one third of the potentiation, and an inhibitory site. Glutamate, GABA and glycine receptors also possess multiple classes of Zn2+ modulatory sites (Paoletti et al., 2000; Laube et al., 2002; Hosie et al., 2003).
Zinc Potentiation Sites Have Structural Similarities to ACh Binding Sites. We compared the zinc potentiation site with other binding sites on Cys-loop receptors. The ACh binding site on nAChRs has been extensively characterized (Corringer et al., 2000). Glu59 is located within the “D-loop” region of α4. In an ACh-binding interface, the D-loop is supplied by the γ, δ, or ϵ subunits of muscle nAChRs or the β subunits of neuronal nAChRs and contributes a critical tryptophan residue to the binding site, as well as several determinants of pharmacological diversity (Corringer et al., 2000; Celie et al., 2004). His162 is located within the “F-loop” region of α4. At ACh binding interfaces, this region is also supplied by the “non-α” subunits of muscle and neuronal nAChRs and contains determinants of pharmacological diversity (Corringer et al., 2000). Thus, the Zn2+-binding site that we have identified has structural similarity to the ACh binding site.
Zinc Potentiation Sites Are Not Related to Calcium Potentiation Sites on Neuronal nAChRs. Neuronal nAChRs are also potentiated by extracellular Ca2+ (Mulle et al., 1992; Vernino et al., 1992). Critical glutamate residues mediating Ca2+ potentiation have been identified on α7 (Galzi et al., 1996). These residues are highly conserved among neuronal nAChRs and correspond to α4Glu45/β4Glu49 (located between β strands 1 and 2) and α4Glu175/β4Glu179 (located just before β strand 9). Both putative Ca2+ binding sites are located at the bottom (near the membrane) of the receptor structure shown in Fig. 1B, away from the Zn2+ potentiation site we have identified. In addition, Ca2+ potentiation of α4β4 is unaffected by the α4H162G and α4E59A mutations (data not shown). Thus, Zn2+ and Ca2+ act at separate sites on neuronal nAChRs.
Zinc Potentiation Sites Are Not Related to Zinc Binding Sites of Other Ligand-Gated Ion Channels. Other Cys-loop receptors also possess Zn2+-binding sites. On the glycine receptor α1 subunit, Asp80 participates in a Zn2+-potentiation site at subunit-subunit interfaces near the top (distal from the cell surface) of the receptor extracellular domain, whereas His107, His109, and Thr112 participate in an inhibitory site that faces into the vestibule (Laube et al., 2002). The Zn2+ site that we have identified on neuronal nAChRs is not structurally comparable with either of these sites. GABAA receptors also possess multiple Zn2+-binding sites. For clarity, it is important to remember that GABAA receptor β subunits are equivalent to neuronal nAChR α subunits, whereas GABAA receptor α subunits are equivalent to neuronal nAChR β subunits. Thus, whereas neuronal nAChRs bind ACh at α-β interfaces, GABA receptors bind GABA at β-α interfaces (Smith and Olsen, 1995). In the α1β3 GABAA receptor, α1Glu137, α1His141 and β3Glu182 participate in a Zn2+-binding inhibitory site located at α-β interfaces (non-GABA binding interfaces) near the bottom (proximal to the cell surface) of the receptor extracellular domain (Hosie et al., 2003). A separate class of Zn2+-binding inhibitory site is located within the ion channel. The Zn2+-potentiation site we have identified on neuronal nAChRs is not structurally comparable with either of these Zn2+-binding sites on GABAA receptors. Glutamate-gated ion channels also possess zinc-binding sites, which have been characterized in detail (Paoletti et al., 2000). However, these receptors are structurally unrelated to nAChRs, precluding a comparison with the Zn2+-potentiation site on neuronal nAChRs.
Zinc Potentiation Sites Are Analogous to the Benzodiazepine-Binding Site on GABAA Receptors. The most interesting comparison is with the binding site for benzodiazepines on GABAA receptors. Whereas GABA binds at β-α interfaces, incorporation of a γ subunit allows benzodiazepines to bind at the α-γ interface (Smith and Olsen, 1995). A series of residues on the γ2 subunit (Tyr58, Phe77, Ala79, and Thr81) have been identified as components of the benzodiazepine-binding site (Kucken et al., 2000, 2003; Teissere and Czajkowski, 2001). Examination of aligned amino acid sequences (Le Novere and Changeux, 2001) and a model of the α1β2γ2 GABAA receptor extracellular domain (Kucken et al., 2003) reveals that the benzodiazepine-binding site residue γ2Thr81 (Teissere and Czajkowski, 2001) is in a location identical to that of α4Glu59. Although α4Glu59 may not directly coordinate zinc, it is close to the zinc potentiation site on neuronal nAChRs. This suggests that the binding of agonists and modulators at alternating subunit-subunit interfaces is a general property of heteromeric Cys-loop receptors. The similar location of the zinc potentiation sites on neuronal nAChRs and the benzodiazepine binding site on GABAA receptors, as well as the similar modulatory function of both sites, leads us to suggest that the Zn2+ potentiation sites on neuronal nAChRs are structurally and functionally similar to the benzodiazepine binding site on GABAA receptors. This identifies the Zn2+ potentiation site on neuronal nAChRs as a promising target for future drug development.
Footnotes
- Received June 9, 2005.
- Accepted September 27, 2005.
This work was supported by National Institutes of Health grants MH66038 and DA08102 (to C.W.L.) and GM069972 (to A.M.). A.M. was supported in part by award BM030 from the Florida Biomedical Research Foundation, and by American Heart Association, Florida/Puerto Rico Affiliate grant SDG-0130456B. B.H., K.B.M., D.E., and S.E.R. were supported in part by National Institutes of Health grant T32-HL07188. B.H. was supported in part by a PhRMA Foundation Medical Student Research Fellowship and is a Lois Pope LIFE Fellow.
B.H and K.B.M. contributed equally to this work.
Article, publication date, and citation information can be found at http://molpharm.aspetjournals.org.
doi:10.1124/mol.105.015164.
ABBREVIATIONS: nAChR, nicotinic acetylcholine receptor; AChBP, acetylcholine binding protein; SCAM, substituted cysteine accessibility method; MTSEA, N-biotinoylaminoethyl methanethiosulfonate; MTSET, [2-(trimethylammonium)ethyl] methanethiosulfonate; ACh, acetylcholine; MTS, methanethiosulfonate.
- The American Society for Pharmacology and Experimental Therapeutics