GABAA receptors, along with the receptors for acetylcholine, glycine, and serotonin, are members of a ligand-gated ion channel superfamily (Ortells and Lunt, 1995). Because of the paucity of crystallographic information for these ligand-gated channels, little is known about the structure of their binding sites or how agonist binding is transduced into channel gating. We used the substituted cysteine accessibility method to obtain secondary structural information about the GABA binding site and to systematically identify residues that line its surface. Each residue from α1 Y59 to K70 was mutated to cysteine and expressed with wild-type β2 subunits inXenopus oocytes or HEK 293 cells. The sulfhydryl-specific reagent N-biotinylaminoethyl methanethiosulfonate (MTSEA-Biotin) was used to covalently modify the cysteine-substituted residues. Receptors with cysteines substituted at positions α1 T60, D62, F64, R66, and S68 reacted with MTSEA-Biotin, and α1 F64C, R66C, and S68C were protected from reaction by agonist. We conclude that α1 F64, R66, and S68 line part of the GABA binding site. The alternating pattern of accessibility of consecutive engineered cysteines to reaction with MTSEA-Biotin indicates that the region from α1 Y59 to S68 is a β-strand.
- GABAA receptor
- binding site
- substituted cysteine accessibility method
- cysteine mutagenesis
- molecular model
GABA is the major inhibitory neurotransmitter in the mammalian brain, and GABAAreceptors are the primary transducers of its action. GABAAreceptors are likely to be heteropentameric proteins (Nayeem, 1994) assembled from distinct subunit classes with multiple subtypes, α(1–6), β(1–4), γ(1–3), δ(1), ε(1), and π(1) (Rabow et al., 1995; Sieghart, 1995; Davies et al., 1997; Barnard et al., 1998). The binding of GABA to GABAA receptors promotes conformational changes leading to the opening of an integral anion-selective channel. Because the GABA binding sites reside on the extracellular surface of the protein and the channel gate is located close to the cytoplasmic end of the channel (Xu and Akabas, 1996), the local changes that occur at the binding site when GABA binds must be propagated to distant parts of the receptor. To understand the transduction of GABA binding to channel gating, one must identify the amino acid residues involved in GABA binding and then locate these residues in a three-dimensional structure of the receptor.
Photoaffinity labeling (Smith and Olsen, 1994) experiments have identified α1F64 as forming part of the GABA binding site. In the β2 subunit, mutations of Y157, T160, T202, and Y205 decrease the apparent affinity of GABA, and the results suggest that these residues are also part of the GABA binding site (Amin and Weiss, 1993). Although mutagenesis and photoaffinity labeling experiments are very useful, these methods cannot identify all the residues that form a ligand binding site or provide detailed structural information about the site.
To systematically identify residues that line the surface of the GABA binding site and to investigate the secondary structure of peptide segments involved in the formation of this site, we used the substituted cysteine accessibility method (Karlin and Akabas, 1998). This approach has been used to identify residues that line the ion-conducting pores of numerous channel proteins (Akabas et al., 1994;Cheung and Akabas, 1996; Kuner et al., 1996; Perez-Garcia et al., 1996;Sun et al., 1996; Xu and Akabas, 1996; Egan et al., 1998) as well as residues forming the surface of the binding site crevice of the dopamine D2 receptor (Javitch et al., 1995; Javitch, 1998).
In the current study, each GABAA receptor residue in the region from α1Y59 to K70 was mutated to cysteine. This region of the receptor was selected for study because it contains α1F64, a binding site residue identified by photoaffinity labeling. Mutant α1 subunits were heterologously expressed with wild-type β2 subunits. A sulfhydryl-specific reagent, N-biotinylaminoethyl methanethiosulfonate (MTSEA-Biotin; Toronto Research Chemicals), was used to covalently modify the substituted cysteines. We identify an engineered cysteine as being in the binding site by two criteria: (1) the reaction with MTSEA-Biotin covalently alters function, and (2) the sulfhydryl-specific reaction is impeded by the presence of binding site ligands.
Here, we show that five residues, α1T60, D62, F64, R66, and S68, are accessible to MTSEA-Biotin. We confirm that α1F64 is part of the GABA binding site, and identify two new binding site residues, α1R66 and S68. By examining the pattern of accessibility of consecutive engineered cysteines, we infer that the region from α1Y59 to S68 is a β-strand. On the basis of these results, a structural model of the GABA binding site is discussed. Because GABA binding is not diffusion-limited and binding most likely depends on receptor structure (Jones et al., 1998), this study provides insight into receptor mechanisms that define GABA affinity.
MATERIALS AND METHODS
Site-directed mutagenesis. The α1cysteine mutant constructs were made either by “Altered Sites II:in vitro Mutagenesis Systems” (Promega, Madison, WI) or by recombinant PCR. Cysteine substitutions were made in the α1 subunit at positions Y59, T60, I61, D62, V63, F64, F65, R66, Q67, S68, W69, and K70 (see Fig. 1). The α1cysteine-substituted mutants were subcloned into pGH19 (Liman et al., 1992; Robertson et al., 1996) for expression in Xenopus laevis oocytes or into pCEP4 (Invitrogen, San Diego, CA) for transient expression in human embryonic kidney (HEK) 293 cells.
All α1 cysteine mutants were verified by double-stranded DNA sequencing. The α1 cysteine mutants have been named using the single letter code, as (wild-type residue) (residue number) (mutated residue).
Expression in oocytes. Oocytes from Xenopus laevis were prepared and injected with cRNA as described previously (Boileau et al., 1998). GABAA receptor rat α1, β2, and α1cysteine mutants in pGH19 were expressed by injection of cRNA into oocytes at molar ratios of 1:1, α/β. The oocytes were maintained in ND96 (in mm): 96 NaCl, 2 KCl, 1 MgCl2, 5 HEPES, 1.8 CaCl2, pH 7.4, supplemented with 100 μg/ml gentamicin and 100 μg/ml BSA for 2–14 d and used for electrophysiological recordings.
Voltage-clamp analysis. Oocytes under two-electrode voltage-clamp (V hold = −80 mV) were perfused continuously with ND96 recording solution at a rate of 5 ml/min. Drugs and reagents were dissolved in ND96. To correct for slow drift in responsiveness, GABA dose–response plots were scaled to a low, nondesensitizing concentration of drug applied just before the drug concentration tested. Standard two-electrode voltage-clamp recording was performed using a GeneClamp 500 (Axon Instruments) interfaced to a computer with an IT-16 A/D device (Instrutech). Electrodes were filled with 3 m KCl and had a resistance of 0.5–1.5 MΩ.
All oocytes were tested for stability of responses to GABA before addition of MTSEA-Biotin by applying two to five pulses of GABA over a period of 10–30 min. The criterion for acceptable stability was that the peak currents varied by <3%. Routinely, GABA concentrations ranged between EC20 and EC60 and were chosen to obtain 0.5–8 μA of current. In general, we tested the covalent effects of MTSEA-Biotin by the following protocol: we determined the peak current evoked by several 5–10 sec applications of GABA, washed for 5 min, applied 2 mm MTSEA-Biotin for 2 min, washed for 5 min, and again determined the peak current evoked by GABA at the same concentration used before MTSEA-Biotin treatment. The covalent effect of MTSEA-Biotin was taken as 1 − (I GABA, after/I GABA, before). To determine whether this response was reversible and repeatable, some cells were incubated for 2 min with ∼20 mm DTT after MTSEA-Biotin exposure and current measurement. After a 15–20 min wash with ND96, current recovery was measured, and in some cases, inhibition by MTSEA-Biotin was tested by repeat exposure.
The protocol for agonist protection experiments was as follows. Various concentrations of MTSEA-Biotin were applied to mutant receptors to determine a low concentration that would yield near-maximal blockage with a 30 sec application. For α1F64C and α1R66C, 50 μm MTSEA-Biotin was chosen; for α1S68C, 200 μm sulfhydryl reagent was required. The effect of those concentrations on mutant receptors served as controls for GABA protection experiments in separate cells. Cells were incubated for 30 sec with the appropriate concentration of MTSEA-Biotin plus a concentration of GABA approximately three times the concentration required for maximal current response (see Fig. 2). After determining the extent of protection from inhibition, the same cells were reexposed to the same concentration of MTSEA-Biotin alone to demonstrate that the full inhibitory effect, as compared with control cells, was obtainable.
Data acquisition and analysis were performed using AxoData, AxoGraph (Axon Instruments), and Prism software (Graphpad). Dose–response data were fit to the following four-parameter equation derived from the Hill equation: Y = Min + (Max − Min)/(1 + 10(LogEC50 -X) · (nH )), where Max is the maximal response, Min is the response at the lowest drug concentration tested, X is the logarithm of agonist concentration, EC50 is the half-maximal response, and nH is the Hill coefficient.
Transient expression in HEK 293 cells. Wild-type rat α1, β2, and cysteine mutant α1 cDNAs in the mammalian expression vector pCEP4 were used for transient transfection of HEK 293 cells (ATCC CRL 1573). Cells were grown on 100 mm tissue culture dishes in Minimum Essential Medium with Earle’s salts (Life Technologies, Gaithersburg, MD) containing 10% fetal bovine serum (Hyclone Laboratories, Logan, UT) in a 37°C incubator under a 5% CO2 atmosphere. Cells were cotransfected at 40–50% confluency with pCEP-α1 or pCEP- α1 cysteine mutant and pCEP-β2. The vector pAdVAntage (Promega) was also added to enhance expression levels (6 μg of each subunit DNA/plate and 12 μg of pAdVAntage). Transient transfection of HEK 293 cells was performed using a standard CaHPO4 precipitation method (Graham and vander Eb, 1973). Cells were harvested, and membrane homogenates were prepared 48 hr after transfection.
Binding assays. Cells were scraped from the dishes and pelleted by centrifugation (1000 × g, 10 min, 4°C). The cells were washed once and resuspended in a HEPES buffer containing (in mm): 124 NaCl, 2.9 KCl, 1.3 MgSO4, 1.2 KH2PO4, 25.0 HEPES, 5.2d-glucose, 2 EDTA, pH 7.4, and homogenized using a Brinkman polytron. The homogenates were centrifuged (30,000 ×g, 20 min, 4°C), and the resulting pellets were resuspended in HEPES buffer. Protein concentrations were determined using a Bradford Assay (Bio-Rad, Hercules, CA) using bovine serum albumin as a standard.
Saturation and competition binding experiments were performed as described previously (Boileau et al., 1998). In brief, membrane homogenates (100 μg) were incubated at room temperature with [3H]muscimol (20 Ci/mmol; DuPont NEN, Wilmington, DE) in a final volume of 250 μl. Nonspecific binding was determined in the presence of 1 mm GABA or 100 μmmuscimol, and specific binding was defined as the amount of tritiated drug bound in the absence of displacing ligand minus the amount bound in the presence of displacer. For saturation binding experiments,K D and B max were determined by fitting specific binding data to a single site using the equation y = (B max *x)/(K D + x), wherey is the specifically bound dpm and x is radiolabeled drug concentration (Prism software; Graphpad). Data from competition binding experiments were fit by using the equationy = B max/(1 + (x/IC50)), where y is the specifically bound dpm, B max is maximal binding, and x is concentration of displacing drug (Prism software; Graphpad). K I was calculated according to the Cheng–Prusoff/Chou equation (Cheng and Prusoff, 1973; Chou, 1974).
MTSEA-Biotin reaction and protection assay in HEK cells. HEK cells were harvested and washed by centrifugation as described above. After the second 1000 × g centrifugation, the cells were gently resuspended in a small volume of HEPES buffer and incubated for 10 min at room temperature with 5 mm MTSEA-Biotin (Toronto Research Chemicals) or buffer as a control. After the incubations, 50 ml of cold HEPES buffer was added, and the cell suspension was centrifuged (2000 × g, 10 min, 4°C). The cells were washed with an additional 50 ml of HEPES buffer, centrifuged (2000 × g, 10 min, 4°C), and then resuspended, and a membrane homogenate was prepared as described above. For protection experiments, cells were incubated for 15 min with 3 mm muscimol (∼50 ×K D) before the incubation with MTSEA-Biotin, and the muscimol remained present during the subsequent incubation with MTSEA-Biotin.
Statistics. We analyzed the effects of MTSEA-Biotin by one-way ANOVA, applying the Dunnett post-test for significance of differences between the effects of MTSEA-Biotin on a mutant receptor and the effects on wild-type receptor (p < 0.01).
Expression of cysteine-substituted receptors inXenopus oocytes
Twelve cysteine mutants were made in the α1 subunit at positions Y59, T60, I61, D62, V63, F64, F65, R66, Q67, S68, W69, and K70 (Fig. 1). Because we test whether an engineered cysteine reacts with MTSEA-Biotin by whether MTSEA-Biotin covalently alters the GABA-induced current in oocytes expressing the mutant, we require that the cysteine substitution mutants be functional. Cysteine mutant α1 subunits were individually expressed with wild-type β2 subunits in Xenopus laevis oocytes, and current responses to GABA were measured. Because expression of single α1 or β2subunits (Boileau et al., 1998) does not produce detectable GABA-mediated chloride currents, a robust current response confirms the expression of both subunits in a fully assembled functional receptor. Application of GABA to receptors containing α1T60C, I61C, D62C, V63C, F64C, F65C, R66C, S68C, and K70C gave robust current responses, whereas no significant GABA-mediated chloride current was detected after expression of α1Q67Cβ2 and α1W69Cβ2 receptors. Thus, cysteine was a functionally tolerated substitute for every residue except α1 Q67 and W69, and it is likely that the positions occupied by the cysteine side chains in the functional mutant receptors are similar to the positions of the native amino acid side chains. Cysteine substitution had little effect (<3.5-fold) on the EC50 for GABA of α1T60Cβ2, α1I61Cβ2, α1D62Cβ2, α1V63Cβ2, α1F65β2, α1S68Cβ2, and α1K70Cβ2 receptors, whereas two mutants, α1 F64Cβ2 and R66Cβ2, had 75-fold and 320-fold increases in EC50, respectively (Table 1, Fig.2).
Reactions of the cysteine-substituted receptors with MTSEA-Biotin in Xenopus oocytes
A 2 min application of 2 mm MTSEA-Biotin had no effect on the currents recorded from wild-type α1β2 receptors or receptors containing α1 I61C, V63C, F65C, and K70C (Fig.3). The result that MTSEA-Biotin had no effect on wild-type receptors suggests either that the free sulfhydryls in wild-type GABAA receptors are inaccessible to MTSEA-Biotin or that reaction with wild-type cysteines has no effect on the function of the receptor. In either case, the absence of effects on wild-type GABAA receptors allows us to interpret the effects of MTSEA-Biotin on cysteine-substituted mutants as covalent modifications of the introduced cysteine.
MTSEA-Biotin had significant effects on the GABA-evoked currents recorded from receptors containing α1 T60C, D62C, F64C, R66C, and S68C (Figs. 3, 4). In receptors containing α1 D62C, F64C, R66C, and S68C, 2 mm MTSEA-Biotin inhibited the subsequent response to GABA by 21, 93, 95, and 61%, respectively. In receptors containing α1 T60C, MTSEA-Biotin increased the GABA response by 56% (Fig. 3). The inhibition and potentiation of the GABA current by disulfide linking of -SCH2CH2(NH)Biotin to the mutant receptors were reversed by treating the oocytes with the reducing agent dithiothreitol (DTT; 20 mm, 2 min) followed by a 15–20 min wash (Fig. 4). After the DTT treatment, MTSEA-Biotin produced the same effect on the GABA-evoked current as before the DTT treatment, demonstrating that the reversibility was caused by reduction of the disulfide bond rather than an artifact of the DTT treatment (Fig. 4).
To determine whether GABA could protect the cysteine mutant receptors from covalent modification by MTSEA-Biotin, a saturating concentration of GABA was added during the sulfhydryl reaction. In these experiments, the duration of the MTSEA-Biotin reaction and its concentration were adjusted so that the minimal amount of MTSEA-Biotin needed to produce a near-maximal effect was used (Figs. 4,5). In the presence of GABA, the reaction of MTSEA-Biotin with receptors containing α1 F64C, R66C, and S68C was significantly inhibited (Figs. 4, 5), whereas the reaction of MTSEA-Biotin with receptors containing α1 T60C and D62C was not changed (data not shown). The presence of GABA caused a 60–70% protection of α1F64Cβ2, α1R66Cβ2, and α1S68Cβ2 receptors, where % protection = (1 − (InhibitionGABA +MTS/InhibitionMTS)) × 100. Because the reaction with MTSEA-Biotin is covalent and the binding of GABA is reversible, complete protection was not observed. Nevertheless, the results indicate that α1 F64, R66, and S68 are near or part of the GABA binding site.
MTSEA-Biotin (2 mm, 2 min) had markedly different magnitudes of effect on some substituted cysteines than on others (Fig.3). MTSEA-Biotin had the largest effects on cysteines substituted for α1 F64 and R66. For cysteines substituted for α1 T60 and S68, MTSEA-Biotin had intermediate effects, whereas MTSEA-Biotin had the smallest effect on α1D62Cβ2 receptors. Even brief exposure to micromolar concentrations of MTSEA-Biotin resulted in almost complete inhibition of current responses of α1F64Cβ2and α1R66Cβ2 receptors (Figs. 4, 5).
Expression of cysteine mutant receptors in HEK 293 cells
To provide additional evidence that the effect of covalently adding -SCH2CH2(NH)Biotin to some of the cysteine-substituted receptors is caused by a direct effect at the binding site, we expressed some of the substituted cysteine α1 subunits with wild-type β2 subunits in HEK 293 cells and examined the ability of MTSEA-Biotin to alter the binding of [3H]muscimol (a GABA agonist) and [3H]SR95531 (a GABA antagonist). Although binding studies with agonists do not necessarily measure binding affinity because agonists induce conformational changes that lead to receptor gating (Colquhoun, 1998), binding studies with antagonists avoid this complication and most likely measure binding directly.
Receptors containing α1 Y59C T60C, I61C, D62C, V63C, F65C, R66C, and S68C specifically bound [3H]muscimol (75–92 nm). At five positions, cysteine substitution had little effect on the affinity of [3H]muscimol binding: α1Y59Cβ2, α1T60Cβ2, α1I61Cβ2, α1V63Cβ2, and α1S68Cβ2 receptors had equilibrium dissociation constants (K D) for [3H]muscimol not significantly different from wild-type α1β2 receptors (Table2). The largest change measured was for α1Y59Cβ2 receptors, which had a 2.9-fold decrease in muscimol affinity as compared with α1β2 receptors. Although specific [3H]muscimol binding was detectable in receptors containing α1 D62C, F64C, F65C, and R66C, the amount of binding was low, and these mutant receptors were not analyzed further. No significant specific [3H]muscimol binding was detected after expression of single α1 or β2 subunits or α1Q67Cβ2 and α1W69Cβ2 receptors.
Reactions of cysteine mutant receptors with MTSEA-Biotin in HEK 293 cells
Cysteine mutant receptors with near-normal binding affinity and expression were analyzed further by covalently reacting them with MTSEA-Biotin. Incubation with MTSEA-Biotin (2 mm, 15 min) caused a 42 ± 2.3% (n = 10) inhibition of [3H]muscimol binding to α1S68Cβ2 receptors and a 40 ± 12% (n = 5) potentiation of binding to α1T60Cβ2 receptors (Fig.6). The binding of [3H]SR95531 (a GABA antagonist) to α1 S68C-containing receptors was also decreased 40% after MTSEA-Biotin treatment (n = 2). MTSEA-Biotin did not have a significant effect on [3H]muscimol binding to α1β2, α1Y59Cβ2, α1I61Cβ2, or α1V63Cβ2 receptors (Fig. 6).
To determine whether muscimol could protect α1T60Cβ2 and α1S68β2 receptors from covalent modification by MTSEA-Biotin, nonradioactive muscimol (3 mm, ∼50 × K D) was added before and during the MTSEA-Biotin reaction. The inhibition caused by the reaction of MTSEA-Biotin with α1S68Cβ2 receptors was 10.3 ± 6% (n = 4) when 3 mm muscimol was added before and during the MTSEA-Biotin reaction (Fig.7). Thus, the presence of 3 mm muscimol caused a 76% protection of α1S68Cβ2 receptors, where % protection = (1 − (10.3/42)) × 100. Addition of 3 mm muscimol to α1T60Cβ2receptors before and during the MTSEA-Biotin reaction did not significantly decrease the potentiation observed (Fig. 7). The results obtained in HEK 293 cells confirm and supplement the data obtained electrophysiologically in Xenopus oocytes and show that α1 S68 is near or part of the GABA binding site.
Residues accessible to MTSEA-Biotin
We used the substituted cysteine accessibility method to investigate the secondary structure of a 12 amino acid segment of the α1 polypeptide chain surrounding F64, a known GABA binding site residue (Sigel et al., 1992; Smith and Olsen, 1994). Furthermore, the approach was used to identify additional residues within this segment that are part of the GABA binding site. We made the following assumptions. (1) The GABA binding site is most likely at a water-accessible surface of the protein because under physiological conditions GABA is zwitterionic (Krogsgaard-Larsen et al., 1984); (2) MTSEA-Biotin is relatively impermeant and reacts preferentially at the water-accessible surface of a protein (Chen et al., 1998); and (3) if a cysteine-substituted residue is part of the GABA binding site, the addition of -SCH2CH2(NH)Biotin will covalently alter binding, and site-selective ligands will protect the introduced cysteine from reaction with MTSEA-Biotin. On the basis of these assumptions, we show that five residues, α1T60, D62, F64, R66, and S68, are solvent-exposed and accessible to MTSEA-Biotin, and three of them, α1F64, R66, and S68, are part of or close to the GABA binding site because GABA slows their reaction with MTSEA-Biotin. Two residues, α1Q67 and W69, do not tolerate cysteine substitution. These two residues are invariant in GABA α (Fig. 1), β, and γ subunits and are highly conserved in all superfamily subunits. We speculate that they play an essential structural role in this receptor superfamily.
The effects of covalently adding -SCH2CH2(NH)Biotin to a substituted cysteine could be attributed to a direct effect such as steric block and/or an indirect allosteric effect on the binding site. Regardless of the mechanism, the observation of a change in receptor function after MTSEA-Biotin treatment is proof that the reaction has occurred. For α1T60Cβ2 receptors, the effect of MTSEA-Biotin is not caused by steric overlap because modification of α1T60C with MTSEA-Biotin leads to a potentiation of both the GABA current response and [3H]muscimol binding (Figs. 4, 6). Furthermore, agonist does not protect α1T60C from MTSEA-Biotin reaction (Fig. 7). For this residue, an indirect effect of the modification leading to an increase in GABA affinity and an enhancement of efficacy (“gating”) are the most likely explanations. MTSEA-Biotin modification of α1D62C decreases GABA-gated current (Fig. 3). This result is consistent with either a direct steric block or an indirect allosteric effect. The fact that GABA does not protect α1D62C from MTSEA-Biotin modification supports an indirect action. However, it is also possible that -SCH2CH2(NH)Biotin, when attached to α1D62C, is long enough to swing into the GABA binding site and sterically hinder GABA binding, although GABA is too small to protect α1D62C from modification. Experiments using sulfhydryl reagents of different sizes will help distinguish between these possibilities. For α1F64Cβ2, α1R66Cβ2, and α1S68Cβ2 receptors, the inhibition measured after MTSEA-Biotin modification and the ability of agonist to protect these residues from modification (Figs. 4, 5, 7) strongly suggest that steric hindrance underlies the inhibition and is consistent with the idea that these residues are near or part of the GABA binding site.
Residues exposed in the GABA binding site
Although allosteric effects cannot be completely ruled out, several lines of evidence argue that α1F64, R66, and S68 line part of the GABA binding site. Results from photoaffinity labeling (Smith and Olsen, 1994) and mutagenesis (Sigel et al., 1992) studies provide evidence that α1F64 is a GABA binding site residue. Our results, showing that the reaction of MTSEA-Biotin with α1F64Cβ2 receptors irreversibly inhibits GABA-mediated chloride current (Fig. 4) and that GABA protects α1F64C from the reaction (Fig. 5), provide independent evidence that α1F64 is part of the binding site. These results demonstrate the validity of using the substituted cysteine accessibility method to identify binding site residues. Thus, on the basis of our criteria and the results reported in this paper, we reason that α1R66 and S68 are also part of or near the GABA binding site.
Further proof that α1F64 and R66 are both in the binding site is provided by the result that introducing cysteines at these positions causes 75- and 320-fold shifts in GABA EC50values of α1F64Cβ2 and α1R66Cβ2 receptors, respectively (Table 1). The shifts in EC50 values are larger than one would predict if the mutations only affected gating (Amin and Weiss, 1993, their Fig. 3b). It is possible, however, that the mutations affect both binding and gating. Interestingly, treatment of purified GABAA receptors with an arginine-specific reagent, 2,3-butanedione, results in a time- and concentration-dependent loss of [3H]muscimol binding (Widdows et al., 1987) and provides supplementary evidence that an arginine residue is important for GABA binding. The ability of MTSEA-Biotin to inhibit not only the GABA-activated chloride current but also the radioligand binding of both a GABA agonist and antagonist to α1S68Cβ2 receptors lends further support for the conclusion that α1S68C is located near the GABA binding site. Finally, the identification of α1R66 and S68 as binding site residues is concordant with their proximity to α1F64 in a β-strand (Fig.8).
Together, these observations are explained most simply by a model in which α1F64, R66, and S68 line part of the GABA binding site (Fig. 8). However, not every one of these residues need to contact GABA. Some of these residues may be important for maintaining the local physico-chemical properties of the site or be involved in the local changes that occur at the binding site when agonist binds. GABA could protect noncontact residues in the binding pocket by blocking the passage of MTSEA-Biotin from the extracellular medium to that particular part of the binding site.
Secondary structure of the polypeptide chain flanking α1F64
Our results, that alternating residues in the primary amino acid sequence from α1Y59 to α1S68 are accessible to MTSEA-Biotin, are consistent with this region forming a β-strand (Fig. 8). Because the accessibility of α1Q67C and α1W69C could not be tested (α1Q67C and α1W69C do not assemble into functional channels), the strictly alternating exposure surrounding α1S68 is not absolutely established. The residues accessible to MTSEA-Biotin, with the exception of α1F64, are hydrophilic amino acid residues. Because MTSEA-Biotin is relatively impermeant (Chen et al., 1998), the accessibility of these residues to reaction suggests that they are exposed at the protein, water-accessible surface. The inaccessible residues are mostly hydrophobic residues and are likely to be buried within the protein. We must be cautious, however, in our interpretation of apparently unreactive residues because we cannot rule out silent reactions that appear to have no functional consequences. Nevertheless, taken together, the results of this study strongly suggest that the polypeptide chain from α1Y59 to S68 forms a β-strand and that a portion of this strand lines the GABA binding site. In agreement with our experimental results, a part of this region (α1 M57-R66) is predicted by secondary structure modeling algorithms (Chou and Fasman, 1978; Smith and Olsen, 1995), to adopt a β-strand conformation.
Theoretical model of the GABA binding site
By analogy to the agonist binding site of the nicotinic acetylcholine receptor (Czajkowski et al., 1993), the GABA binding site of the GABAA receptor has been proposed to lie at the interface between the α and β subunits (Galzi and Changeux, 1994;Smith and Olsen, 1995). We propose that one domain of the GABA binding site on the α1 subunit is formed in part by a β-strand and that α1F64, R66, and S68 are facing into the GABA binding site (Fig. 8). Previous mutagenesis studies (Amin and Weiss, 1993) have suggested that two domains on the β2 subunit, Y157 -T160 and T202-Y205, also form part of the GABA binding site. Although experimental evidence is lacking, we have tentatively modeled these segments as two α-helices because the identified residues in each segment are three residues apart. We are currently using the substituted cysteine accessibility method on these β2subunit domains to directly test this hypothesis.
The orientation of GABA relative to these identified binding site residues is not known. The stabilization of GABA binding will most likely involve electrostatic interactions and hydrogen bonding between GABA’s charged groups and the side chains of binding site amino acid residues. We speculate that an electrostatic interaction between the positive guanidinium group of α1R66 and the negative carboxyl group of GABA stabilizes GABA binding. α1R66 is conserved in all GABAA receptor α, π, and ρ subunits. At the positive end of GABA, hydrogen bonding with β2T160 and Y157 as well as interactions with the aromatic ring of α1F64 may be important. Experiments using engineered GABA affinity reagents that can be “tethered” to cysteines substituted for α1F64, R66, or S68 will be helpful in determining GABA’s exact placement in the site.
These studies are a step toward constructing a detailed molecular model of the GABA binding site and ultimately will help explain how GABA binds and initiates the conformational changes that result in anion channel opening. Because the GABA binding site is most likely formed by residues from two adjacent subunits, we hypothesize that GABA and other agonists bridge the binding site. Agonist binding could promote a change in the distance between the α1 and β2 subunits that causes a shift of one subunit relative to the other, and this movement could then be propagated to the opening of the channel. With the methods described in this report and sulfhydryl-specific cross-linking reagents, we are now in a position to test this and alternative hypotheses.
C.C. is a recipient of the Burroughs Welcome Fund New Investigator Award in the Basic Pharmacological Sciences. This work was supported in part by National Institute of Neurological Diseases and Stroke Grant NS34727 to C.C. We thank Dr. Jean-Yves Sgro for expert assistance with the molecular modeling, Dr. Nicholas Cozzi and Allison Friedlein for help during the initial stages of this project, Amy Kucken for technical assistance, Drs. Meyer Jackson, Larry Trussell, and David Wagner for critical reading of this manuscript, and Dr. Jonathan Javitch for invaluable discussions.
Correspondence should be addressed to Dr. Cynthia Czajkowski, University of Wisconsin, Department of Physiology, Room 197 MSC, 1300 University Avenue, Madison, WI 53706.