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The Journal of Neuroscience, February 15, 2006, 26(7):2031-2040; doi:10.1523/JNEUROSCI.4555-05.2006

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Cellular/Molecular
Charged Residues in the ₁ and ₂ Pre-M1 Regions Involved in GABA_A Receptor Activation

Jose Mercado and Cynthia Czajkowski

Department of Physiology and Neuroscience Training Program, University of Wisconsin–Madison, Madison, Wisconsin 53706

	Abstract

Top Abstract Introduction Materials and Methods Results Discussion References

 
For Cys-loop ligand-gated ion channels (LGIC), the protein movements that couple neurotransmitter binding to channel gating are not well known. The pre-M1 region, which links the extracellular agonist-binding domain to the channel-containing transmembrane domain, is in an ideal position to transduce binding site movements to gating movements. A cluster of cationic residues in this region is observed in all LGIC subunits, and in particular, an arginine residue is absolutely conserved. We mutated charged pre-M1 residues in the GABA_A receptor ₁ (K219, R220, K221) and ₂ (K213, K215, R216) subunits to cysteine and expressed the mutant subunits with wild-type ₂ or ₁ in Xenopus oocytes. Cysteine substitution of ₂R216 abolished channel gating by GABA without altering the binding of the GABA agonist [³H]muscimol, indicating that this residue plays a key role in coupling GABA binding to gating. Tethering thiol-reactive methanethiosulfonate (MTS) reagents onto ₁K219C, ₂K213C, and ₂K215C increased maximal GABA-activated currents, suggesting that structural perturbations of the pre-M1 regions affect channel gating. GABA altered the rates of sulfhydryl modification of ₁K219C, ₂K213C, and ₂K215C, indicating that the pre-M1 regions move in response to channel activation. A positively charged MTS reagent modified ₂K213C and ₂K215C significantly faster than a negatively charged reagent, and GABA activation eliminated modification of ₂K215C by the negatively charged reagent. Overall, the data indicate that the pre-M1 region is part of the structural machinery coupling GABA binding to gating and that the transduction of binding site movements to channel movements is mediated, in part, by electrostatic interactions.

Key words: GABA; GABA_A receptor; pre-M1 region; substituted cysteine accessibility method; electrostatic interactions; allosteric protein; ligand-gated ion channel

	Introduction

Top Abstract Introduction Materials and Methods Results Discussion References

 
For ligand-gated ion channels (LGIC), neurotransmitter binding triggers a series of conformational movements that ultimately lead to channel opening. Because the neurotransmitter binding site resides on the extracellular surface of the protein and the channel gate is located in the transmembrane domain (TMD), the local changes that occur at the binding site when neurotransmitter binds must be propagated to distant parts of the receptor protein. Although much is known about the structure of the Cys-loop superfamily of LGICs from the recent 4 Å resolution model of the Torpedo nicotinic acetylcholine receptor (nAChR) (Unwin, 2005) and the crystallographic structure of the related acetylcholine binding protein (AChBP) (Brejc et al., 2001; Celie et al., 2004), these structures alone cannot describe the protein dynamics involved in coupling neurotransmitter binding to channel gating.

The N-terminal region of the first transmembrane domain (pre-M1), a region that structurally links the extracellular ligand binding domain (LBD) to the transmembrane channel domain, is in an ideal position to transduce binding site movements in the extracellular domain to gating movements in the membrane domains. It physically links -strand 10, which forms part of loop C of the binding site, to the first transmembrane -helix M1. In addition, a cluster of cationic residues in this region is observed in all LGIC subunits, and in particular, the arginine at ₂216 of the GABA_A receptor (GABA_AR) and aligned positions is absolutely conserved (see Fig. 1), suggesting that charge interactions may play a role in coupling ligand binding to channel gating. Here, we used the substituted cysteine accessibility method to examine the electrostatic environment and dynamics of the pre-M1 region of the GABA_A receptor during channel activation.

Charged amino acid residues in the pre-M1 regions of the ₁ and ₂ subunits of the GABA_A receptor were individually mutated to cysteine. The rates these cysteines were modified by sulfhydryl-specific methanethiosulfonate (MTS) reagents were measured in unliganded (resting) and GABA-bound receptor states (open/desensitized) to determine whether structural movements occur in the pre-M1 region during channel activation. In addition, we used MTS reagents of different charge to monitor the electrostatic environment near the pre-M1 region.

We demonstrate that both the ₁ and ₂ pre-M1 regions undergo structural rearrangements during channel activation, as expected if this region is an important transduction element coupling binding to gating. Moreover, we provide evidence that mutation of the conserved arginine residue at position 216 in the ₂ subunit to cysteine uncouples agonist binding from gating. Oocytes expressing ₁₂R216C receptors had no functional response to GABA but specifically bound the GABA agonist [³H]muscimol to similar levels as wild-type receptors. Differences in the accessibility and the functional effects of oppositely charged MTS compounds suggest that unliganded and liganded receptor state stabilization is mediated, in part, by electrostatic forces. Overall, the data indicate that the pre-M1 region is part of the structural machinery linking binding site movements to channel gating movements.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results Discussion References

 
Mutagenesis. Rat cDNAs encoding ₁ and ₂ GABA_A receptor subunits were used for all molecular cloning and functional studies. Cysteine mutants were made using either the QuickChange site-directed mutagenesis kit (Stratagene, La Jolla, CA) or by recombinant PCR, as described previously (Boileau et al., 1999). Mutagenic oligonucleotides were synthesized to introduce the desired mutation, and a silent restriction site was used to screen for the desired mutations. All mutant cDNA was verified by double-stranded cDNA sequencing to confirm that the desired point mutations was present and that the cDNA was free of additional mutations.

Expression in Xenopus laevis oocytes. Oocytes were prepared as described previously (Boileau et al., 1998). Capped cRNAs encoding the ₁, ₂, ₁K219C, ₁R220C, ₁K221C, ₂K213C, ₂K215C, and ₂R216C subunits in the vector pGH19 (Liman et al., 1992; Robertson et al., 1996) were transcribed in vitro using the mMessage mMachine T7 kit (Ambion, Austin, TX). Single oocytes were injected within 24 h with 27 nl of cRNA (10 ng/µl per subunit) in a ratio of 1:1. Oocytes were incubated at 18°C in ND96 (in mM: 96 NaCl, 2 KCl, 1 MgCl₂, 1.8 CaCl₂, and 5 HEPES, pH 7.2), supplemented with 100 µg/ml gentamycin and 100 µg/ml BSA, for 2–14 d before use.

Two-electrode voltage clamp. Oocytes were perfused continuously at a rate of 5 ml/min with ND96 while being held under two-electrode voltage clamp at –80 mV. The bath volume was 200 µl. Stock solutions of GABA (Sigma, St. Louis, MO), muscimol (Sigma), and pentobarbital (Research Biochemicals, Natick, MA) were prepared fresh in ND96. Borosilicate electrodes (Warner Instruments, Hamden, CT) were filled with 3 M KCl and had resistances between 0.7 and 2 M. Electrophysiological data were acquired with a GeneClamp 500 (Molecular Devices, Sunnyvale, CA) interfaced to a computer with an ITC16 analog-to-digital device (Instrutech, Great Neck, NY) and recorded using the Whole Cell program, version 3.2.9 (kindly provided by J. Demspter, University of Strathclyde, Glasgow, UK).

Concentration–response analysis. Concentration–response experiments were performed as described previously (Boileau and Czajkowski, 1999). In brief, GABA concentration–responses were scaled to a low, nondesensitizing concentration of GABA (EC₂ to EC₁₀) applied just before the test concentration to correct for any slow drift in I_GABA responsiveness over the course of the experiment. Currents elicited by each test concentration were normalized to the corresponding low concentration current before curve fitting. GraphPad (San Diego, CA) Prism 4 software was used for data analysis and fitting. Concentration–response data were fit to the following equation: I = I_max/(1 + (EC₅₀/[A])ⁿ), where I is the peak response to a given concentration of GABA, I_max is the maximum amplitude of current, [A] is the agonist concentration, and n is the Hill coefficient.

Modification of introduced cysteine residues by MTS reagents. Three derivatives of methanethiosulfonate (CH₃SO₂X) were used to covalently modify the introduced cysteines: MTS-N-biotinylaminoethyl [X = SCH₂CH₂NH-biotin (MTSEA-biotin)], MTS-ethyltrimethylammonium [X = SCH₂CH₂N(CH₃)₃⁺ (MTSET⁺)], and MTS-ethylsulfonate [X = SCH₂CH₂ SO₃^– (MTSES^–)] (Biotium, Hayward, CA). Stocks solutions (100 mM) were made in DMSO for all MTS reagents, aliquoted into microcentrifuge tubes, and rapidly frozen on ice before storage at –20°C. For each application of MTS reagent, a new aliquot was thawed, diluted in ND96 to the working concentration, and used immediately to avoid hydrolysis of the MTS compound. The final DMSO concentrations were 2%. These solvent concentrations did not affect GABA_A receptor functional responses.

MTS modifications of the engineered cysteines were assayed by measuring changes in GABA-evoked current (I_GABA). The effects of MTSEA-biotin, MTSET⁺, and MTSES^– were studied using the following protocol: GABA (EC_40–60) current responses (10–30 s) were measured from oocytes expressing wild-type (₁₂) or mutant receptors and stabilized. Stability was defined as <5% variance of peak current responses to GABA on two consecutive applications. After stabilization, the MTS reagent (2 mM) was bath applied for 2 min, followed by a 5–7 min wash, and then I_GABA was measured at the same concentration as before the MTS treatment. The effect of the MTS application was calculated as follows: [((I_after/I_initial) – 1) x 100], where I_after is the peak GABA current elicited after the MTS application and I_initial is the peak current before MTS. Maximal I_GABA was also measured, before and after MTS application using 10 and 300 mM GABA. In these cases, the time interval between GABA applications was 9–12 min to allow complete recovery from desensitization.

Rate of MTS modification. The rates at which the various MTS reagents modified the engineered cysteines were determined by measuring the effect of sequential applications of low concentrations of MTS reagents on I_GABA, as described previously (Holden and Czajkowski, 2002). The protocol is described as follows: EC_40–60 GABA was applied for 10 s every 3–5 min until I_GABA stabilized (<3% variance). After a 40 s ND96 washout, MTS reagents were applied for 5–20 s, and the cell was then washed for an additional 2.5–4.5 min. The procedure was repeated until I_GABA no longer changed, indicating that the reaction had proceeded to apparent completion. Concentration of MTS reagent and time of application varied as follows: ₁K219C: MTSEA-biotin, 10 µM, 20 s; ₁K221C: MTSEA-biotin, 10 µM, 20 s; MTSES^–, 300 µM, 20 s; ₂K213C: MTSET⁺, 30 µM, 20 s; MTSES^–, 500 µM, 20 s; ₂K215C: MTSET⁺, 30 µM, 20 s; MTSES^–, 300 µM, 20 s. The effects of GABA on the rate of MTS modification were measured by coapplying GABA (EC_80–90) with the MTS reagent. For these experiments, I_GABA was stabilized as follows: EC_40–60 GABA was applied for 10 s, washed for 40 s; EC_80–90 GABA was applied for 5–20 s, washed for 2.5–5 min. The procedure was repeated until I_GABA from EC_40–60 GABA was within <3% of the previous I_GABA peak. This ensured complete washout of the drug and that any alteration in the current amplitudes after MTS treatment was the result of MTS modification. Concentrations of MTS reagents and times of applications in the presence of GABA (EC_80–90) were as follows: ₁K219C: MTSEA-biotin, 30 µM, 20 s; ₁K221C: MTSEA-biotin, 10 µM, 20 s; ₂K213C: MTSET⁺, 30 or 60 µM, 10 s; ₂K215C: MTSET⁺, 30 µM, 10 s.

For all rate experiments, the decrease or increase in GABA-induced current was plotted versus cumulative time of MTS exposure. Peak current at each time point was normalized to the initial peak current (t = 0) and fit to a single exponential function using GraphPad Prism software. A pseudo-first-order rate constant (k₁) was determined, and a second-order rate constant (k₂) was calculated by dividing k₁ by the concentration of the MTS reagent used (Pascual and Karlin, 1998).

[³H]Muscimol single oocyte binding. For binding experiments, the concentration of cRNA injected was 5.4 ng per subunit. Intact oocytes were washed twice with PBS (in mM: 137.93 NaCl, 2.67 KCl, 1.5 KH₂PO₄, and 8 Na₂HPO₄·7H₂O, pH 7.1). To measure total [³H]muscimol binding sites, intact oocytes (8 or 10) were incubated on ice in PBS with 150 nM [³H]muscimol (15.7 Ci/mmol; DuPont NEN, Boston, MA) in triplicate (final volume, 0.5 ml). Nonspecific binding was determined in the presence of 100 µM muscimol. After a 1 h incubation on ice, the samples were vacuum filtered over GF/B glass microfiber filters (Whatman, Clifton, NJ) using a Hoefer Scientific (San Francisco, CA) vacuum box (model FH224) and washed four times with 5 ml of ice-cold PBS. The filters and oocytes were mixed thoroughly with 4 ml of scintillation fluid, and the radioactivity (in counts per minute) was determined. Specific binding was defined as the amount of [³H]muscimol bound in the absence of displacing ligand (total) minus the amount bound in the presence of 100 µM muscimol (nonspecific).

Statistical analysis. Log (EC₅₀) values, changes in GABA EC₅₀ and maximal response after MTS modification, and second-order (k₂) rates were analyzed generally using a one-way ANOVA, followed by a post hoc Dunnett’s test to determine the level of significance between wild-type and mutant receptors. Comparison of the second-order (k₂) rates of covalent modification of ₂K213C and ₂K215C in the absence and presence of GABA were performed using Student’s one-tailed paired t test. Alterations in GABA EC₅₀ resulting from MTS treatment were analyzed using Student’s two-tailed paired t test.

Structural modeling. A model of the extracellular LBD of the GABA_A receptor was built based on the structure of the AChBP (Brejc et al., 2001). The crystal structure of the AChBP was downloaded from Research Collaboratory for Structural Bioinformatics Protein Data Bank (code 1I9B [PDB] ) and loaded into Swiss Protein Bank Viewer (SPDBV; http://www.expasy.org/spdbv/). The rat ₁ mature protein sequence from Thr¹² to Ile²²⁷, the ₂ protein sequence from Ser¹⁰ to Leu²¹⁸, and the ₂ protein sequence from Gly²⁵ to Arg²³¹ were aligned with the AChBP primary amino acid sequence as depicted by Cromer et al. (2002) and threaded onto the AChBP tertiary structure using the "Interactive Magic Fit" function of SPDBV. The threaded subunits were imported into SYBYL (Tripos, St. Louis, MO), where energy minimization was performed. SYBYL minimizations terminate when the number of iterations is reached or if the gradient change reaches 0.05 kcal/Å·mol. The first 100 iterations were performed using Simplex minimization (Press et al., 1998) followed by 1000 iterations using the Powell conjugate gradient method (Powell, 1977). A GABA_A receptor LBD (:::: viewed counterclockwise from the synaptic cleft) was assembled by overlaying the monomeric subunits on the AChBP scaffold, and the resulting structure was imported into SYBYL and energy minimized. Neither water nor entropy factors were included during the minimizations. After the global energy minimization, Ramachandran plots, plots, side-chain positions, and cis and trans bonds were all examined. The program PROCHECK (Laskowski et al., 1996) was run to examine structural features against the established database of protein parameters, most importantly the / torsions and side-chain conformations. Problems in the structure that were revealed by these evaluations were fixed manually, and energy minimizations were run again as needed. Regions with insertions were modeled by fitting structures from a loop database. Because the sequence identity of the AChBP and the GABA_A receptor LBD is only 18%, caution must be used in interpreting the absolute positions of individual side-chain residues in the model.

The structure of the nAChR transmembrane domain at 4 Å resolution solved by Miyazawa et al. (2003) (PDB 1OED [PDB] ) was used to model the pore-containing TMDs of the GABA_A receptor. The GABA_A receptor TMD sequences were aligned with the nAChR TMD sequences and the amino acids from the ₁, ₂, and ₂ subunits of the GABA_AR were substituted for residues in the /, , and nAChR subunits, respectively. The TMD of each subunit was built and energy minimized individually using the Tripos force field program in SYBYL. A GABA_A receptor TMD pentamer was assembled by overlaying the monomeric subunits on the nAChR scaffold, and the resulting structure was imported into SYBYL and energy minimized as described above.

A model of the entire GABA_A receptor (LBD plus TMD) was built by physically docking the LBD model with the TMD model using the fivefold axes as a docking guide. The domains were rotated so that their relative positions to one another were consistent with published experimental data. The TMD N-terminal and the LBD C-terminal residues were joined and energy minimized while fixing the rest of the molecule. This was followed by an energy minimization of the entire structure in three stages. The first stage used the Simplex method for 20 cycles before switching to the Tripos force field for 1000 iterations. Finally, the third stage again used the Tripos force field with >1000 cycles, or a convergence of <0.05 kcal/Å·mol. After the global energy minimization, Ramachandran plots, plots, side-chain positions, and cis and trans bonds were all examined. The program PROCHECK (Laskowski et al., 1996) was run to examine structural features against the established database of protein parameters, most importantly the / torsions and side-chain conformations. Problems in the structure that were revealed by these evaluations were fixed manually and energy minimizations were run again as needed. Although the resulting model is reasonable, the packing of the loops at the interface of the LBD and TMD as well as the positions of residue side-chains need to be interpreted with care.

	Results

Top Abstract Introduction Materials and Methods Results Discussion References

 
Functional characterization of pre-M1 mutant receptors
Three charged residues in the ₁ subunit (K219, R220, and K221) and in the ₂ subunit (K213, K215, and R216) were mutated to cysteine in the pre-M1 region (Fig. 1). Oocytes were injected with each of the mutant subunits, wild-type ₂, or ₁ and were assayed for GABA responsiveness using two-electrode voltage clamp. GABA elicited currents (I_GABA) from oocytes expressing ₁K219C₂, ₁K221C₂, ₁₂K213C, and ₁₂K215C receptors, indicating that these cysteine substitutions were tolerated and yielded functional receptors. The GABA EC₅₀ and Hill coefficient values were not significantly different from wild-type values (EC₅₀ = 6.9 ± 0.7 µM; n_H = 1.0 ± 0.1) (Fig. 2, Table 1). Oocytes expressing ₁K219C₂, ₁K221C₂, ₁₂K213C, and ₁₂K215C receptors elicited maximal macroscopic currents similar to wild-type receptors (2130 ± 885 nA; n = 8).

View larger version (28K): [in this window] [in a new window]   Figure 1. Aligned amino acid sequences of the pre-M1 and adjacent regions of the rat (R) GABAA receptor 1, 2, and 2 subunits, rat glycine receptor 1 subunit, rat serotonin type-3 receptor subunit, and the T. californica (T.c.) nicotinic acetylcholine receptor 1 subunit. Aligning GABAA receptor subunit sequences with other LGIC subunits reveals a cluster of charged residues in the pre-M1 region. Cationic and anionic residues in the pre-M1 regions are in boldface. The GABAA receptor arginine residues at 1220 and 2216 are conserved in all LGIC subunits (underlined). Residues mutated to cysteine are marked with a "C" above the residue. Residues located in the GABA binding site, 2Y205 and 2R207 (Amin and Weiss, 1993; Wagner and Czajkowski, 2001), and the binding sites of other LGICs (Kao et al., 1984; Mishina et al., 1985; Dennis et al., 1988; Ruiz-Gomez et al., 1990; Galzi et al., 1991a,b; Vandenberg et al., 1992a,b; Rajendra et al., 1995) are marked with an asterisk. Elements of secondary structure are indicated below the sequences by the arrow (-strand 10) and rectangle ( helix, M1 transmembrane region).

 

View larger version (25K): [in this window] [in a new window]   Figure 2. GABA concentration–response curves of wild-type 12 and mutant GABAA receptors. A, Representative current responses from an oocyte expressing 12K213C receptors elicited by increasing concentrations of GABA (micromolar). B, GABA concentration–response curves from oocytes expressing 12 (; dashed line), 1K219C2 (), 1K221C2 (), 12K213C (), and 12K215C () receptors. Data points represent the mean ± SEM from three to four independent experiments. Data were fit by nonlinear regression analysis as described in Materials and Methods. GABA EC50 and nH values are reported in Table 1.

 

View this table: [in this window] [in a new window]   Table 1. GABA EC50 and Hill coefficient values of wild-type and mutant receptors

 
Neither GABA (1 mM) nor pentobarbital (1 mM) elicited current responses from oocytes expressing ₁R220C plus ₂ or ₁ plus ₂R216C combinations of subunits (Fig. 3A), suggesting that cysteine substitutions at these positions affected receptor assembly/surface expression or that surface mutant receptors were expressed but were unresponsive to GABA and pentobarbital. To help distinguish between these possibilities, we measured the binding of [³H]muscimol to surface receptor protein using an intact oocyte binding assay. Intact oocytes expressing ₁₂R216C receptors specifically bound similar amounts of [³H]muscimol (4 ± 0.9 fmols per oocyte; n = 3) as wild-type ₁₂ receptors (3 ± 0.4 fmols per oocyte; n = 3) (Fig. 3B), suggesting that the ₂R216C mutation does not appear to affect receptor expression or agonist binding but uncouples ligand binding from channel gating. Oocytes expressing ₁R220C₂ receptors did not bind [³H]muscimol and were not gated by GABA or pentobarbital (Fig. 3), suggesting that cysteine substitution of ₁R220 was not tolerated.

View larger version (13K): [in this window] [in a new window]   Figure 3. Expression of 12, 1R220C2, and 12R216C GABAA receptors. A, Representative GABA- (1 mM) and pentobarbital- (PB; 1 mM) mediated current responses from oocytes injected with 12 [wild type (WT)], 1R220C plus 2, and 1 plus 2R216C cRNA. Neither GABA nor pentobarbital elicited current responses from oocytes expressing 1R220C or 2R216C subunits. B, Specific [3H]muscimol binding was measured in intact oocytes expressing WT and mutant subunits as described in Materials and Methods. Data were normalized to the specific [3H]muscimol binding measured for WT receptors. Noninjected oocytes (N.I.) and oocytes injected with 1R220C2 subunits did not specifically bind [3H]muscimol. Oocytes expressing 12R216C receptors specifically bound similar amounts (129 ± 28%) of [3H]muscimol as oocytes expressing WT receptors. Error bars represent the mean ± SEM from three independent experiments.

 
MTS effects on substituted cysteines
To explore the physicochemical environment of the pre-M1 region, we examined the ability of MTS reagents that differ in size and charge to modify wild-type, ₁K219C₂, ₁K221C₂, ₁₂K213C, and ₁₂K215C receptors. The MTS reagents (Fig. 4A) used were: MTSEA-biotin, which covalently adds a neutral biotinylaminoethyl group (12 Å long); MTSET⁺, which adds a positively charged ethyl-trimethylammonium group (4.5 Å long); and MTSES^–, which adds a negatively charged ethyl-sulfonate group (4.8 Å long). Application of 2 mM MTSEA-biotin, MTSET⁺, and MTSES^– for 2 min to wild-type receptors had no effect on I_GABA (EC₅₀ concentration), indicating that any effects observed in the mutant receptors were caused by covalent modification of the introduced cysteines (Fig. 4B). MTSEA-biotin modification of ₁K219C, ₂K213C, and ₂K215C increased I_GABA by 34 ± 2, 34 ± 9, and 34 ± 2% (n 3), respectively, and decreased I_GABA by 60 ± 4% (n = 9) in ₁K221C-containing receptors (Fig. 4B).

View larger version (38K): [in this window] [in a new window]   Figure 4. Effects of MTS reagents on wild-type and mutant GABAA receptors. A, Structures of MTS reagents represent the portions of the reagents that covalently modify an introduced cysteine. Lengths were measured after energy minimization (<0.5 kcal/Å; ChemSketch; Advanced Chemistry Development, Toronto, Ontario, Canada). B, Effects of a 2 min application of 2 mM MTSEA-biotin, MTSET+, or MTSES– on wild-type (WT) and mutant receptors. The percentage change in IGABA after MTS treatment is defined as follows: [((Iafter/Iinitial) – 1) x 100]. Negative values represent an inhibition of IGABA after MTS reaction, whereas positive values represent an increase in IGABA. Data represent the mean from at least four independent experiments. All of the effects observed on the mutated receptors after MTS modification, except for 1K221C2 plus MTSET+ and 1K219C2 plus MTSES–, were statistically significant from wild type (p < 0.01).

 
Derivatization of ₂K213C and ₂K215C with MTSES^– increased I_GABA by 47 ± 5 and 48 ± 7%, respectively, whereas MTSET⁺ increased I_GABA by 95 ± 8 and 53 ± 4%, respectively (n 8) (Fig. 4B). In contrast, modification of ₁K219C₂ and ₁K221C₂ by MTSES^– and MTSET⁺ differentially affected I_GABA (Fig. 4B). Tethering a negative charge (MTSES^–) on ₁K221C₂ enhanced I_GABA (98 ± 14%; n = 4), whereas treatment with the positively charged MTSET⁺ had no effect on I_GABA (Fig. 4B). To determine whether MTSET⁺ reacted with ₁K221C, we first applied MTSET⁺ and then MTSES^– to the same oocyte. The ability of MTSES^– to potentiate subsequent GABA currents was eliminated by pretreatment with MTSET⁺ (data not shown), indicating that MTSET⁺ covalently modified ₁K221C, but modification had no effect in our functional assay (silent reaction). In contrast to the effects seen on ₁K221C, tethering MTSET⁺ on ₁K219C decreased I_GABA (31 ± 7%; n = 3), whereas modification with MTSES^– had no effect (silent reaction). The different functional effects measured after MTSES^– and MTSET⁺ modification of ₁K219C and ₁K221C are likely caused by differences in the local electrostatic environments near these residues rather than steric effects, because MTSET⁺ and MTSES^– are similar in size (Fig. 4A) and have a common reaction mechanism.

As described above, cysteine substitution of ₂R216 abolished channel gating by GABA without altering [³H]muscimol binding (Fig. 3). To determine whether restoring the positive charge at this position would restore channel gating by GABA, we measured whether GABA would elicit current responses from ₁₂R216 receptors after MTSET⁺ exposure. Application of 2 mM MTSET⁺ for 2 min to ₁₂R216C receptors had no effect on I_GABA (data not shown), suggesting that tethering the positively charged ethyl-trimethylammonium group from MTSET⁺ was insufficient to restore GABA_AR current responses or that the introduced cysteine was not modified.

MTS effects on maximal GABA current
An increase or decrease in I_GABA after MTS application can be attributed to a change in GABA apparent affinity (EC₅₀) and/or a change in maximal GABA response (I_max). To test whether MTS modification altered GABA I_max, we measured current responses elicited from GABA EC₅₀ concentrations (I_EC50) and saturating concentrations of GABA (10 and 300 mM; I_max) before and after MTS application (Fig. 5). MTSEA-biotin was used for the ₁ mutants (₁K219C₂, ₁K221C₂) and MTSET⁺ for the ₂ mutants (₁₂K213C, ₁₂K215C). Application of 2 mM MTSEA-biotin or MTSET⁺ for 2 min to wild-type receptors did not affect I_EC50 or I_max (Fig. 5). MTS modification significantly increased I_max for ₁K219C₂ (34 ± 3%), ₁₂K213C (37 ± 7%), and ₁₂K215C (43 ± 5%) receptors (n 4; p < 0.01), suggesting that tethering thiol-reactive groups at these positions caused changes in channel gating or conductance. MTSEA-biotin significantly reduced I_max by 55 ± 3% (Fig. 5) for ₁K221C₂ receptors (n = 7; p < 0.01). Although the results are consistent with a decrease in channel gating at this position, we cannot rule out the possibilities that a subset of receptor channels might be blocked by the MTS compound or that modification causes an enormous increase in the unbinding of GABA, which could contribute to the decrease in I_max measured.

View larger version (18K): [in this window] [in a new window]   Figure 5. Effects of MTS reagents on GABA maximal currents. A, Representative GABA-mediated current traces from oocytes expressing wild-type (12) and 12K213C receptors. Currents elicited by an EC50 concentration of GABA and a maximum concentration of GABA (10 mM; ECmax) were recorded before and after a 2 min MTS (2 mM) exposure. For 12K213C receptors, MTSET+ modification potentiated both GABA EC50 and ECmax responses. B, Summary of the peak current responses elicited by GABA ECmax (filled circles) and GABA EC40–60 (open circles) concentrations before (–) and after (+) MTS application for 12 and mutant receptors are shown. Data for each receptor were normalized to the peak response to GABA ECmax before (–) addition of the MTS reagent and represent the mean ± SEM from at least three independent experiments. MTSET+ and MTSEA-biotin (MTSB) treatment had no effect on 12 GABAA receptor current responses but significantly altered the current responses elicited by GABA EC50 and ECmax concentrations (p < 0.01) for all mutant receptors.

 
To estimate whether the MTS modifications were also altering GABA apparent affinity, we calculated the I_EC50/I_max ratio of current responses (R) after MTS treatment for each mutant receptor. Assuming that MTS modification did not significantly alter the Hill coefficients, we would expect a ratio of 0.5 if the MTS modification had no effect on GABA EC₅₀, a ratio >0.5 if GABA EC₅₀ decreased, and a ratio <0.5 if GABA EC₅₀ increased. The I_EC50/I_max ratio did not alter significantly for ₁K219C₂ (R_before, 0.44 ± 0.02; R_after, 0.43 ± 0.02) and ₁₂K215C receptors (R_before, 0.47 ± 0.02; R_after, 0.50 ± 0.03) but increased significantly for ₁₂K213C receptors (R_before, 0.47 ± 0.04; R_after, 0.67 ± 0.05 µM) and decreased significantly at ₁K221C₂ receptors (R_before, 0.46 ± 0.02; R_after, 0.29 ± 0.05 µM) (n 3; p < 0.05), suggesting that covalent modification of ₁K221C and ₂K213C altered GABA apparent affinity. No apparent changes in macroscopic desensitization after MTS treatment were observed.

MTSET⁺ and MTSES^– reaction rates
The electrostatic potential near an introduced cysteine can be measured by comparing the second-order reaction rates of a negatively and a positively charged MTS reagent (Karlin and Akabas, 1998). For ₁₂K213C and ₁₂K215C receptors, MTSET⁺ rates were 80 and 43 times faster than MTSES^– rates, respectively (Fig. 6, Table 2). In contrast, the rate of MTSET⁺ modification of the simple thiol 2-mercaptoethanol in solution is only 12.5-fold faster than the rate with MTSES^– (Table 2). To factor out the intrinsic differences in the reactivity of the two reagents, we divided the ratio of the rates of the two reagents at an introduced cysteine by the ratio of the rates for the two reagents with mercaptoethanol (Stauffer and Karlin, 1994; Cheung and Akabas, 1997; Yang et al., 1997). For ₂K213C, the ratio of ratios is = 80/12.5 = 6.4 and for ₂K215C, = 43/12.5 = 3.4. A ratio of ratios of 1 indicates that there is no charge selectivity at that position. A ratio of ratios that is significantly larger than one indicates that there is a negative potential experienced by the thiol.

View larger version (21K): [in this window] [in a new window]   Figure 6. MTSET+ and MTSES– reaction rates. A, Representative current traces recorded while measuring the rate of MTSET+ modification of 12K213C receptors in the absence of GABA. GABA EC40–60 current responses were recorded before and after successive application (10–20 s) of 30 µM MTSET+ (arrows). B, Normalized GABA current responses were plotted versus cumulative time of MTSET+ () or MTSES– (500 µM; ) exposure and fit with single-exponential functions. Data were normalized to the current measured at time = 0 and are (mean ± SEM) from at least four independent experiments. Second-order rate constants (k2) are reported in Table 2.

 

View this table: [in this window] [in a new window]   Table 2. Second-order rate constants for MTSES– and MTSET+ modification of 12K213C and 12K215C receptors

 
We can estimate the effective electrostatic potential at an introduced cysteine by using the following equation: = –(1/z_MTSET – z_MTSES)(RT/F)ln(), where z is the unitary charge of the MTS reagent, R is the gas constant, T is absolute temperature, F is Faraday’s constant, and is the ratio of ratios calculated above (Stauffer and Karlin, 1994; Yang et al., 1997; Karlin and Akabas, 1998). The calculated negative electrostatic potentials at ₂K213C and ₂K215C in the resting state are –24 and –15 mV, respectively.

Effect of GABA on MTS reaction rates
To determine whether the pre-M1 regions undergoes structural rearrangements during channel gating, we measured the rates of MTS modification of ₁K219C, ₁K221C, ₂K213C, and ₂K215C in the absence ("closed," resting state) and presence of near-saturating GABA (open/desensitized states). The closed reaction rates were all 2000 M^–1·s^–1 (Fig. 7D, Table 3). For ₁₂K213C and ₁₂K215C receptors, the MTSET⁺ reaction rates were three and four times faster in the presence of EC_80–90 GABA, respectively (Fig. 7, Table 3). In contrast, MTSEA-biotin modified ₁K219C approximately three times slower than in its absence, and GABA had no significant effect on the rate of modification of ₁K221C (Fig. 7C, Table 3). The alterations in the rates of modifications of ₁K219C, ₂K213C, and ₂K215C in the presence of GABA demonstrate that GABA activation of the receptor triggers movements in or near the pre-M1 regions of both the ₁ and ₂ subunits.

View larger version (19K): [in this window] [in a new window]   Figure 7. Rates of MTS modification of 1 and 2 pre-M1 mutant GABAA receptors in the presence and absence of GABA. A, Rate of sulfhydryl modification of 12K215C receptors in the absence and presence of GABA. Representative current traces recorded while applying MTSET+ (30 µM) in the absence (top) and presence (bottom) of GABA (100 µM; EC90). GABA EC40–60 current responses were recorded before and after successive applications (10–20 s) of 30 µM MTSET+ alone or coapplied with GABA (arrows). B, Normalized GABA current responses were plotted versus cumulative time of MTSET+ () and MTSET+ coapplied with EC80–90 GABA () and fit with single-exponential functions. Data were normalized to the current measured at time = 0 and represent mean ± SEM from at least three independent experiments. C, Summary of the second order rate constants (k2) for reaction of MTS reagents with mutant receptors in the absence (MTS) and presence of GABA (+GABA). Second-order rate constant (k2) are reported in Table 3. **Values significantly different from MTS alone (p < 0.001).

 

View this table: [in this window] [in a new window]   Table 3. Second-order rate constants (k2) for reaction of MTS reagents with mutant receptors in the absence (control) and presence of GABA

 
Recently, it has been proposed that ₂D146 (loop 7, Cys–Cys loop) moves closer to ₂K215C (pre-M1) in the open/desensitized state (Kash et al., 2004). If a negatively charged residue is moving closer to the pre-M1 region during receptor activation, then this could explain the increases in the rates of MTSET⁺ modification of ₂K213C and ₂K215C measured when GABA is present (Fig. 7, Table 3). To test this hypothesis, we examined the ability of MTSES^– (2 mM) to modify ₂K215C in the presence of GABA. We reasoned if GABA activation of the receptor resulted in an increase in the negative electrostatic potential near ₂K215C, then reaction with MTSES^– would be slowed or eliminated in the presence of GABA. Application of MTSES^– alone increased I_GABA 47 ± 5% (Fig. 4). Interestingly, when GABA was coapplied with MTSES^–, the treatment had no effect. To determine whether MTSES^– reacted silently with ₂K215C, we applied MTSES^– first in the presence of GABA and then in its absence (data not shown). When MTSES^– was subsequently applied in the absence of GABA, GABA currents were potentiated, indicating that MTSES^– does not covalently modify ₂K215 when the receptor is bound with GABA. The data are consistent with the idea that activation of the receptor by GABA induces an increase in the negative electrostatic potential near ₂K215C, which inhibits MTSES^– from reacting with the introduced cysteine.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

 
Structural pathway linking agonist binding to channel gating
Structural studies suggest that acetylcholine binding to the nAChR induces a 10° clockwise rotation of the inner strands of the extracellular ligand binding domain of the subunit. It is postulated that this rigid body rotation is then translated to the transmembrane M2 channel-lining helix via the ₁–₂ loop (loop 2) (Miyazawa et al., 2003; Unwin, 2005). Functional studies, using rate–equilibrium free energy relationships, suggest a temporal sequence of events in which agonist binding triggers a conformational wave that begins at the binding site, followed by movements of loops 2 and 7 (Cys–Cys loop) and then the M2–M3 linker, and finally movements of the transmembrane helices (Grosman et al., 2000a,b; Chakrapani et al., 2004). A recent study using a chimeric receptor comprised of AChBP fused to the transmembrane pore domain of the serotonin type-3A (5-HT_3A) receptor showed that only when loops 2, 7, and 9 from AChBP were replaced with 5-HT_3A receptor sequence was functional communication between the ligand binding site and channel domain restored, indicating that these loops are critical elements involved in coupling ligand binding to channel gating (Bouzat et al., 2004). It should be noted, however, that the above chimeric receptor contained the charged pre-M1 region of the 5-HT_3A receptor, which raises the possibility that the pre-M1 region is also involved in transducing binding to gating.

Here, we provide evidence that the pre-M1 regions of the GABA_A receptor ₁ and ₂ subunits play pivotal roles in coupling binding site movements to the channel domain. The rates of MTS modification of ₁K219C, ₂K213C, and ₂K215C were significantly altered in the presence of GABA (Fig. 7, Table 3), demonstrating that the pre-M1 regions undergo structural rearrangements in response to channel activation, as predicted if the pre-M1 regions propagate structural movements from the binding site to the transmembrane domain. Tethering thiol-reactive groups onto these same residues (₁K219C, ₂K213C, and ₂K215C) increased maximal GABA-activated currents (Fig. 5), suggesting that structural perturbations of the pre-M1 regions alter channel gating. Finally, cysteine substitution of ₂R216 abolished channel gating by GABA without altering [³H]muscimol binding (Fig. 3), indicating that this residue plays a key role in allosterically coupling GABA binding to gating. Interestingly, the ₂R216C mutation also abolished channel gating by pentobarbital and suggests that this residue may be part of a common activation pathway used by both drugs.

Additional support for a role of the ₂ subunit pre-M1 region in transducing binding site movements to channel gating movements comes from its location. It physically links loop C of the binding site to the membrane domain. Structural studies (Unwin et al., 2002; Celie et al., 2004) as well as molecular dynamic simulations (Henchman et al., 2005) reveal that agonist binding promotes movement of loop C inward to cap the binding site. We predict that this capping movement is propagated to the channel transmembrane domain by the pre-M1 region. In particular, ₂R216 appears to be a key player. This residue is absolutely conserved in every member of the Cys-loop receptor superfamily, and its mutation has been shown to alter agonist-induced channel gating in other Cys-loop receptors (Vicente-Agullo et al., 2001; Hu et al., 2003; Castaldo et al., 2004). A model of the GABA_A receptor shows ₂R216 in close proximity to ₂E52 (Fig. 8), which is also highly conserved, suggesting that these residues electrostatically interact and potentially form a salt bridge linking the pre-M1 region and loop 2. Using this structural pathway, we envision agonist-induced movements in the loop C region of the binding site being propagated to the pre-M1 region, loop 2, and the top of the M2 channel lining helix, which can then bring about channel opening.

View larger version (46K): [in this window] [in a new window]   Figure 8. Structural model of the GABAA receptor 2 subunit. A, Domains believed to contribute to the transduction mechanism (loop 2, loop 7, M2–M3 linker, and pre-M1) are highlighted in yellow. The pore-forming transmembrane domain 2 (M2) is highlighted in red. Pre-M1 residues 2K213 and 2K215 as well as nearby anionic residues are shown in "stick" format. 2R207, a residue located in the GABA binding site, is shown in a space-filled format. B, Detailed view of the interface between the ligand binding domain and transmembrane domain, which highlights the proximity of residue 2R216 and 2E52, which is located in loop 2.

 
Electrostatic network of interactions
The precise details of how the pre-M1 region influences these movements still remain to be worked out. Although the 4 Å structure of the nicotinic acetylcholine receptor provides information about the path of the peptide backbone, residue side-chain position is difficult to determine at this resolution, especially in regions without secondary structure such as those regions that are likely mediating the interactions between the extracellular ligand binding domain (loop 2, loop 7, loop 9, pre-M1) and the transmembrane domain (extracellular end of M1, extracellular end of M2, M2–M3 loop). Here, we demonstrate that the pre-M1 residues K219C, K221C, K213C, and K215C are located on the water-accessible protein surface, because charged MTS reagents were able to modify these residues (Fig. 4).

Cysteine substitutions of the conserved pre-M1 lysine residues had no effects on GABA EC₅₀ (Fig. 2, Table 1), and thus, the positions occupied by the cysteine side chains in the mutant receptors are likely similar to the native lysine positions. This allowed us to probe the electrostatic environment of the ₂ pre-M1 region using MTS reagents that differ in charge. MTSET⁺ modified ₁₂K213C and ₁₂K215C receptors significantly faster than MTSES^– (Fig. 6, Table 2). In the resting/closed state, we calculated a negative electrostatic potential of –24 mV near ₂K213C and a potential of –15 mV near ₂K215C. A single glutamate residue can contribute approximately –13.5 mV to a local potential (Stauffer and Karlin, 1994). Consistent with this data, in a homology model of the GABA_A receptor, two negatively charged residues ₂D190 (on -strand 9) and ₂E147 (loop 7) are in close proximity to ₂K213, whereas ₂D146 (loop 7) is near ₂K215 (Fig. 8). The electrostatic potential near the ₁ pre-M1 region could not be measured, because MTSES^– had no functional effect on ₁K219C and MTSET⁺ had no effect on ₁K221C (Fig. 4). However, the model reveals ₁D148 (loop 7) in close proximity to ₁K219; ₁D54 (loop 2), ₁E143 (loop 7), and ₁D144 (loop 7) near ₁R220; and ₁D191 (on -strand 9) near ₁K221C (Fig. 9).

View larger version (49K): [in this window] [in a new window]   Figure 9. Structural model of the GABAA receptor 1 subunit. A, Domains believed to contribute to the transduction mechanism (loop 2, loop 7, M2–M3 linker, and pre-M1) are highlighted in yellow. The pore forming transmembrane domain 2 (M2) is highlighted in red. Pre-M1 residues 1K221 and 1K219 as well as nearby anionic residues are shown in "stick" format. B, Detailed view of the interface between the ligand binding domain and transmembrane domain, which highlights two anionic residues (1E143 and 1D144) located in loop 7 nearby the conserved residue 1R220 at the pre-M1 region.

 
Previous studies have suggested that charged residues in loops 2, 7, and the M2–M3 loop play important roles in LGIC activation (Grosman et al., 2000a,b; Absalom et al., 2003; Kash et al., 2003, 2004; Bouzat et al., 2004; Mukhtasimova et al., 2005). It has been reported that an electrostatic interaction between ₂D146 (loop 7) and ₂K215 (pre-M1) is critical for receptor activation based on a double charge swap mutation (₂D146K, K215D) restoring GABA EC₅₀ to near wild-type values compared with single charge reversals (Kash et al., 2004). If gating is controlled by the pair-wise interaction between these residues, one would predict that cysteine mutation of ₂K215, which removes the charge, should alter GABA EC₅₀. This was not observed (Fig. 2, Table 1). Notably, interaction energies obtained from double mutant cycles reflect not only direct pair-wise interactions but can also reflect structural perturbations of neighboring residues as well as interactions between networks of nearby residues (Schreiber and Fersht, 1995), particularly if a residue is coupled to others. Our data demonstrating that structurally perturbing the pre-M1 region influences channel gating (Fig. 5) and the finding that significant coupling energies are measured between ₂K215D and a variety of receptor mutations including ₂D139K, ₂D146K, ₂E52K, ₂D56K, and ₂E147K [Kash et al. (2004), their Tables 1, 2] are consistent with ₂K215 and the pre-M1 region being part of a larger network of interacting residues.

Thus, we envision that a network of interactions between residues in the pre-M1 region, loop 2, loop 7, and the top of M2 exists and that coupling of neurotransmitter binding and channel gating (opening, closing, desensitization, and resensitization) is controlled by the forming and breaking of specific sets of contacts between residues in these regions. As a result of the numerous charged residues located in these regions, many of the interactions are likely to be electrostatic. Our data demonstrating that GABA activation of the receptor results in an increase in the negative electrostatic potentials near ₂K215 and ₂K213 (Table 3) as well as previous data in the GABA_AR ₁ subunit demonstrating interactions between charged residues in loop 2, loop 7, and the M2–M3 loop (Kash et al., 2003) are consistent with this idea. The kinetics of salt bridge breaking is estimated to be 200 ns (Sheldahl and Harvey, 1999; Gruia et al., 2003), and thus, the breaking and forming of salt bridges is potentially a fast enough mechanism for transducing movements from the binding site to the channel domain.

Experiments using rate–equilibrium free energy relationships (Chakrapani et al., 2004) suggest that the transduction of binding to gating occurs as sequential, coupled movements of "rigid body domains," with the ligand binding domain moving first ( 0.9), loop 2 and loop 7 next ( 0.8), then the M2–M3 extracellular loop (