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Originally published In Press as doi:10.1074/jbc.M604752200 on October 26, 2006

J. Biol. Chem., Vol. 281, Issue 51, 39708-39718, December 22, 2006
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Influence of Agonists and Antagonists on the Segmental Motion of Residues near the Agonist Binding Pocket of the Acetylcholine-binding Protein*

Ryan E. Hibbs{ddagger}§, Zoran Radic{ddagger}, Palmer Taylor{ddagger}, and David A. Johnson1

From the {ddagger}Department of Pharmacology and the §Biomedical Sciences Graduate Program, University of California San Diego, La Jolla, California 92093 and the Division of Biomedical Sciences, University of California Riverside, Riverside, Calfornia 92521

Received for publication, May 17, 2006 , and in revised form, October 24, 2006.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Using the Lymnaea acetylcholine-binding protein as a surrogate of the extracellular domain of the nicotinic receptor, we combined site-directed labeling with fluorescence spectroscopy to assess possible linkages between ligand binding and conformational dynamics. Specifically, 2-[(5-fluoresceinyl)aminocarbonyl]ethyl methanethiosulfonate was conjugated to a free cysteine on loop C and to five substituted cysteines at strategic locations in the subunit sequence, and the backbone flexibility around each site of conjugation was measured with time-resolved fluorescence anisotropy. The sites examined were in loop C (Cys-188 using a C187S mutant), in the beta9 strand (T177C), in the beta10 strand (D194C), in the beta8-beta9 loop (N158C and Y164C), and in the beta7 strand (K139C). Conjugated fluorophores at these locations show distinctive anisotropy decay patterns indicating different degrees of segmental fluctuations near the agonist binding pocket. Ligand occupation and decay of anisotropy were assessed for one agonist (epibatidine) and two antagonists ({alpha}-bungarotoxin and d-tubocurarine). The Y164C and Cys-188 conjugates were also investigated with additional agonists (nicotine and carbamylcholine), partial agonists (lobeline and 4-hydroxy,2-methoxy-benzylidene anabaseine), and an antagonist (methyllycaconitine). With the exception of the T177C conjugate, both agonists and antagonists perturbed the backbone flexibility of each site; however, agonist-selective changes were only observed at Y164C in loop F where the agonists and partial agonists increased the range and/or rate of the fast anisotropy decay processes. The results reveal that agonists and antagonists produced distinctive changes in the flexibility of a portion of loop F.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The nicotinic acetylcholine receptor (nAChR)2 represents a group of acetylcholine-gated cation channels that are prototypic of the Cys-loop superfamily of pentameric ligand-gated ion channels that includes the {gamma}-aminobutyric acid, types A and C, 5-HT3 and glycine receptors. nAChRs are primarily responsible for fast neurotransmission in both the peripheral and central nervous systems, and nAChR isoforms are defined by their subunit composition that in turn determines their ligand selectivity, cation permeability, and channel gating kinetics (1).

Before the availability of an atomic-resolution model of the nAChR, the agonist/antagonist binding site was localized to subunit interfaces and mapped with reference to the primary subunit sequences by mutagenesis and chemical modification. Seven segments that appeared to form the agonist/antagonist binding pocket were identified and arbitrarily denoted alphabetically as segments A-C (in the so-called "principal" subunit face that include the distinctive vicinal cysteines in loop C) D-F (in the neighboring and so-called "complementary" subunit) (2, 3), and a final segment involved in binding peptidic toxins (4).

Great insight into the structure of the nAChR has come from analyses of x-ray crystallographic structures of the acetylcholine binding protein (AChBP) (5-9). AChBPs are soluble homopentameric proteins that are found in several salt and fresh water mollusks and share close structural identity with the extracellular domain of the Torpedo nAChR. AChBP appears to be both a structural and functional surrogate for the extracellular domain of the nAChR. In fact, acetylcholine activates a channel of a chimera formed from a modified AChBP and the ion-channel domain of the 5-HT3 receptor (10). This finding indicates that the molecular basis for ligand gating of ion channels is conserved across the entire pentameric ligand-gated ion channel superfamily. The most recent electron micrograph reconstruction of the receptor in Torpedo electroplax membranes has yielded a 4-Å-resolution model of the Torpedo nAChR, and by superimposing the AChBP structure on to the receptor a detailed template of the entire receptor has been developed (11).

A fundamental question in receptor structure-function relationships now focuses on the molecular basis for agonist activation of these channels since the residues that form the extracellular binding site lie some distance from the ion gate in the transmembrane domain. The most studied allosteric gating theory posits a series of intraprincipal subunit rigid-body movements that starts with a twisting, inward movement of the beta9-beta10 hairpin (loop C) toward acetylcholine as it binds in a crevice formed by the beta9-beta10 hairpin and a portion of the beta7-beta8 loop (loop B) in the principal subunit and elements of the beta5 and beta6 strands in the complementary subunit (8, 11-13). This movement is then thought to effect a displacement of the beta1-beta2 loop next to the transmembrane helix (M2) lining the ion channel in the same subunit as the beta9-beta10 hairpin, that in turn leads to channel opening. We reasoned that this intrasubunit mechanistic linkage or one involving the adjacent subunit would require agonist-specific changes in the {alpha}-carbonyl backbone mobility along an activation pathway that extends to the transmembrane ion gate.

To examine the potential role of regions near the acetylcholine binding site in initiating conformational changes that may result in an activation signal, we measured the influence of nicotinic ligands on {alpha}-carbon backbone flexibility using AChBP as a receptor surrogate and a combination of site-directed labeling and fluorescence spectroscopy. Specifically, fluorescein was selectively conjugated to five substituted cysteines and a free cysteine in separate mutants of the Lymnaea stagnalis AChBP, and the backbone flexibility around each site of conjugation was assessed by monitoring the time-resolved fluorescence anisotropy decay of the reporter group. Of the six sites examined, one was in the beta9-beta10 hairpin at the tip of loop C (Cys-188 using a C187S mutant), one in the beta9 strand (T177C), one was in the beta10 strand (D194C), two were in the loop connecting the beta8 and beta9 strands (N158C and Y164C, loop F), and one was in the beta7 strand (K139C). In the absence of ligand regional variation in {alpha}-carbon backbone mobility was observed, with loop C being the most ordered. Ligand-induced changes in loop C mobility did not correlate with agonist-antagonist behavior; however, the loop F site Y164C was found to undergo agonist-specific changes in mobility.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Ligands and Labeling Reagents—(+)Epibatidine, {alpha}-bungarotoxin, nicotine, lobeline, and carbamylcholine were purchased from Sigma-Aldrich. d-Tubocurarine chloride was from ICN Pharmaceuticals, Inc. Methyllycaconitine citrate (MLA) was from Tocris (Ellisville, MO). 4-Hydroxy,2-methoxybenzylidene anabaseine (4-OH,2MeOBA) was obtained from Dr. William Kem at the University of Florida. 2-[(5-Fluoresceinyl)aminocarbonyl]ethyl methanethiosulfonate (MTS-Fl) was purchased from Toronto Research Chemicals, Inc. (Ontario, Canada). {alpha}-125I-Labeled bungarotoxin (specific activity, 130 Ci/mmol) and (+/-)-[3H]epibatidine (specific activity, 65 Ci/mmol) were products of PerkinElmer Life Sciences. Tetramethylrhodamine-{alpha}-bungarotoxin (TMR-Bgt) was purchased from Invitrogen. All other chemicals were of the highest grade commercially available.

Expression, Mutagenesis, and Purification of AChBP—Wild-type AChBP from L. stagnalis was expressed from a cDNA synthesized from oligonucleotides selected for mammalian codon usage, as previously described (14, 15). Briefly, the cDNAs were inserted into a pFLAG-CMV-3 expression vector (Sigma) containing a preprotrypsin leader peptide followed by an NH2-terminal 1x FLAG epitope. A COOH-terminal His6 tag was attached to the protein for radioligand binding assays. Stable cell lines of single cysteine mutants of AChBP were generated as previously described (16). AChBPs, typically in amounts between 4 and 6 mg, were purified from tissue culture medium by adsorption onto an {alpha}-FLAG antibody column and elution with FLAG peptide as previously described (16). Purity and assembly of subunits as a pentamer were assessed by SDSPAGE and fast protein liquid chromatography. We attempted to engineer single cysteines at sites in loop C other than those presented in this study, but our attempts (T184C, Y185C, S186C, P189C, E190C, Y192C, E193C) resulted in non or weak binding, presumably misfolded protein.

Radioligand Binding Assays—A scintillation proximity assay (Amersham Biosciences) was adapted for use in a soluble radioligand binding assay as previously described (16). Briefly, AChBP (0.5 nM binding sites) was incubated with increasing concentrations of either 125I-labeled {alpha}-bungarotoxin or (+/-)-[3H]epibatidine in a solution of 0.1 mg/ml anti-His scintillation proximity assay beads. In competition assays, 125I-labeled {alpha}-bungarotoxin was held constant at 20 nM, and the competing ligand was added in variable concentrations. Radioactivity was measured on a Beckman LS 6500 liquid scintillation counter. Conversions from EC50 to KD were made with the Prism 4 software package from GraphPad Software, Inc (San Diego, CA) using a sigmoidal dose-response plot with a variable slope fitting. All radioligand binding data are averages of at least three replicate experiments.

MTS-Fl Labeling—For each mutant, MTS-Fl and AChBP were dissolved in 100 µl of 50 mM Tris-HCl, 150 mM NaCl, 0.02% NaN3, pH 7.4, to final binding site concentrations of 100 and 20 µM, respectively. After 60 min at room temperature and shielded from light, the reaction mixtures were eluted through G-25 Sephadex columns (20 x 1 cm; Amersham Biosciences) and equilibrated with 0.1 M sodium phosphate buffer, pH 7.0, to remove unconjugated MTS-Fl.

Specific labeling was assessed by comparison of fluorophore emission at equilibrium from the labeled mutant with that of a sample of WT AChBP that was labeled in parallel with the mutant after standardization to protein concentration by UV absorbance. Steady-state emission spectra were measured at room temperature using a Jobin Yvon/Spex FluoroMax II spectrofluorometer (Instrument S.A., Inc., Edison, NJ) with the excitation and emission bandwidths set at 2 nm. In all cases nonspecific labeling was ≤5%. Stoichiometry of labeling for each preparation was estimated from a comparison of fluorophore concentration (absorbance at 496 nm for MTS-Fl, extinction coefficient 85,000 M-1 cm-1) and protein concentration (by absorbance at 280 nm, extinction coefficient 268,000 M-1 cm-1). Stoichiometries of labeling for each mutant were as follows: K139C, 19%; N158C, 10%; Y164C, 32%; T177C, 19%; C187S, 22%; D194C, 37%.

Stopped-flow Kinetic Measurements—Stopped-flow experiments on ligand association and dissociation kinetics were conducted using an Applied Photophysics SX.18MV (Leatherhead, UK) stopped-flow spectrofluorometer. The Fl-Cys-188 AChBP mutant was excited at 490 nm, and a 515-nm cut-on filter was used to select the fluorescence signal. The second-order association rate constants for binding of {alpha}-bungarotoxin and epibatidine were determined from the slope of plots of the observed rate of fluorescence change versus ligand concentration. The first-order rate constant of {alpha}-bungarotoxin dissociation was determined by mixing the preformed AChBP:{alpha}-bungarotoxin complex with concentrations of epibatidine in large excess over its KD and measuring the rate of resulting change in fluorescence. Because binding of epibatidine resulted in a smaller fluorescence enhancement than {alpha}-bungarotoxin, a decrease in fluorescein emission was observed. The first-order rate constant of epibatidine dissociation was measured by mixing the preformed AChBP-epibatidine complex with concentrations of gallamine (which alone gave no fluorescence enhancement) in large excess over its KD and observing the time course of the decrease in fluorescein emission. Equilibrium dissociation constants were determined as a ratio of the dissociation and association rate constants. Similar experiments were performed with Fl-D194C to monitor epibatidine binding and Fl-Y164C to monitor {alpha}-bungarotoxin binding. In the case of Fl-D194C the competing ligand used to determine the dissociation rate was {alpha}-bungarotoxin, and for Fl-Y164C the competing ligand was epibatidine.

Estimation of Ligand Dissociation Constants by Fluorescein to Tetramethylrhodamine Fluorescence Resonance Energy Transfer (FRET)—In cases where ligand binding yielded little change in fluorescein fluorescence, we employed TMR conjugated to {alpha}-bungarotoxin to measure ligand occupation. Proximity of the Fl-TMR pair yields donor (Fl) quenching and acceptor (TMR) fluorescence sensitization. Excitation and emission spectra of this fluorescent pair shows excellent donor-acceptor overlap free from interference from the native protein fluorescence (17). The titrations provide a direct estimate of the dissociation constant for the substituted toxin. Association of competing ligands were estimated by back titration and concomitant reversal of FRET.

Time-resolved Fluorescence Anisotropy—Emission anisotropy was determined as previously described (18). Unless stated otherwise, emission anisotropy decay was analyzed with the impulse reconvolution method implemented in the DAS6TM software package from HORIBA Jobin Yvon IBH Ltd. (Glasgow, UK) described elsewhere (19). Briefly and simply, this approach splits the analysis into two steps, analysis of the total emission decay, S(t), followed by analysis of the vertical/perpendicular difference emission decay, D(t). S(t), free of anisotropy effects, is given by the expression

Formula(Eq.1)

and was analyzed as a biexponential function. D(t), which includes both fluorescence and anisotropy parameters, is given by the expression

Formula(Eq.2)

D(t) is deconvolved with the results from the S(t) analysis as a constraint, yielding

Formula(Eq.3)

D(t) is deconvolved with the results from the S(t) analysis as a constraint, yielding

Here, beta1 and beta2 are the amplitudes of the anisotropy at time 0 for the fast and slow anisotropy decay processes, respectively. {varphi}1 and {varphi}2 are the fast and slow rotational correlation times of the anisotropy decay, respectively. We define the fractional magnitude of the observable anisotropy decay associated with the "fast" diffusional processes as fxb, which is equal to beta1/(beta1 + beta2), and the ratio fxb/{varphi}1, which is a complex function of the rate and range of fast diffusional processes that are usually associated with segmental motion around each site of fluorophore conjugation (20). A nonassociative model was assumed, where the emission relaxation times are common to all the rotational correlation times. Goodness of fit was evaluated from the values of the reduced {chi}2r and by visual inspection of the weighted residual plots.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Production and Characterization of Cysteine Mutants—Six single-residue substitution mutants of the AChBP from Lymnaea were engineered; five cysteine substitution mutants (K139C, N158C, Y164C, T177C, and D194C) and a serine substitution mutant (C187S) (Fig. 1A). The C187S mutant prevented the normal disulfide bond formation between Cys-187 and Cys-188 and, in turn, allowed for selective labeling of Cys-188. Radioligand binding assays were used with a pair of reference ligands for each AChBP mutant to verify that mutagenesis had not affected the overall fold or structure of the binding site (Table 1). Direct saturation binding measurements were made with {alpha}-125I-labeled bungarotoxin, and in separate experiments the KD for epibatidine (Fig. 2) was determined by competition against the radiolabeled {alpha}-neurotoxin. All the mutants retained a high affinity for both ligands assayed. The largest deviations from wild-type affinity observed for the substituted cysteines was the D194C mutant, which lost ~25-fold in binding affinity for {alpha}-bungarotoxin, and the C187S mutant, which lost ~200-fold in binding affinity for epibatidine. Even in these most-extreme cases, the cysteine-substituted AChBP mutants bind the reference ligands with dissociation constants in the low to mid nanomolar range. Therefore, we conclude that the interaction determinants are maintained but with lower interaction energy for the D194C and C187S substitution mutants. Losses in binding affinity at other mutation positions were judged to be small and non-significant when compared with the unmodified enzyme.


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  		TABLE 1
KD values for AChBP mutants with reference ligands

For direct saturation binding, {alpha}-125I-labeled bungarotoxin (I) was incubated in increasing concentrations with AChBP from Lymnaea at 0.5 nM in binding sites. Bound ligand was measured by a scintillation proximity assay (16). For competition experiments epibatidine was incubated in increasing concentrations with AChBP, 0.5 nM in binding sites and a constant concentration of {alpha}-125I-labeled bungarotoxin of 20 nM. Measurements were made in triplicate, and variance from the mean was less than 30%. Conversions from IC50 to KD were made using Prism version 4 (GraphPad Software, Inc.), where KD = IC50/(1 + [I]/KI).

 
Determination of the Ligand Binding Parameters of the Fluorescently Labeled AChBPs—In the case of the C187S mutant devoid of the native vicinal disulfide bond, a significant loss of interaction energy occurred, and we were concerned that at this site in particular, conjugation of the cysteine with MTS-Fl could further reduce ligand binding. We observed, however, that binding of both {alpha}-bungarotoxin and epibatidine to the Fl-labeled mutant resulted in a substantial enhancement of steady-state fluorescence. Accordingly, using stopped-flow measurements of ligand association and dissociation rates, we were able to determine the KD from the ratio of kinetic constants for both {alpha}-bungarotoxin and epibatidine at the covalently modified binding site. Although both {alpha}-bungarotoxin and epibatidine lost approximately 2 orders of magnitude in binding affinity to the Fl-Cys-188 protein, they still bound to AChBP with appreciable affinities (KD = 1.6 and 0.05 µM, respectively), strongly suggesting substantial retention of the configuration of the binding site (Table 2).


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  		TABLE 2
KD values for representative AChBP-fluorescein conjugates

Measurements were made in triplicate, and variance from the mean was less than 30%.

 


Figure 1
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  		FIGURE 1.
Positions of fluorescein labeling of the AChBP and time-resolved fluorescence anisotropy decay for apoAChBP. Panel A, ribbon diagram of the x-ray structure of two adjacent AChBP subunits from Lymnaea (PDB accession code 1I9B (5)), with the principal subunit in blue and the complementary subunit in red. Five-amino acid side chains were mutated to cysteine for subsequent fluorescein labeling (Lys-139, Asn-158, Tyr-164, Thr-177, Asp-194) and one free cysteine was created (Cys-188) by replacing its disulfide bonding partner (Cys-187) with a serine residue. In the orientation presented in this figure, the top of AChBP corresponds to the apical portion of the nAChR, and the bottom is where the transmembrane {alpha}-helices of the nAChR would attach. Panel B, time-resolved fluorescence anisotropy decays of fluorescein-conjugated AChBPs in the absence of ligand. The plot labeled Lamp is the instrument response function.

 


Figure 2
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  		FIGURE 2.
Structures of the fluorescent probe MTS-Fl and the nicotinic ligands used in anisotropy decay analyses. The pharmacologic activities of the majority of ligands used in these experiments have been well studied (38). Epibatidine, nicotine, and carbamylcholine are classical full agonists at muscle and neuronal nAChRs, whereas d-tubocurarine, {alpha}-bungarotoxin, and MLA are competitive antagonists. Lobeline is reported as a mixed agonist/antagonist of nAChRs; in AChBP it induces conformations, mimicking other agonists (8). 4-OH,2MeOBA is the primary active metabolite of 2,4-dimethoxy-benzylidene-anabaseine (also known as GTS-21) and shows partial agonism selective for {alpha}7 receptors (39).

 
For the cysteine substitutions and conjugations that exhibited little or no change in fluorescence anisotropy upon ligand binding, it was necessary to demonstrate that the ligands studied are in fact bound to the Fl-labeled binding site at the concentrations used in the anisotropy experiments. To this end steady-state emission spectra for Fl-T177C were measured at room temperature as described for the determination of nonspecific labeling (See "Experimental Procedures"). Saturation binding of a TMR conjugate of {alpha}-bungarotoxin was monitored upon excitation of fluorescein at 485 nm by observing the quenching of fluorescein fluorescence emission at 515 nm; incremental increases in the concentration of the TMR-Bgt resulted in incremental decreases in Fl-T177C fluorescein emission due to FRET from fluorescein to TMR (Fig. 3, A and B). Because the minimum concentration of binding sites required for a fully quantifiable signal was ~100 nM, the dissociation constant determined in this manner for the fluorescein-labeled T177C mutant could only be estimated to be ≤45 nM (Table 2). By comparison, the dissociation constants for both WT and unlabeled T177C AChBPs were determined by radioligand binding competition experiments with [3H]epibatidine to be 17 nM. Hence, the TMR-labeled toxin has an affinity for AChBP comparable with the native toxin, and fluorescein labeling of the T177C mutant results in at most a 3-fold reduction in affinity for the labeled toxin. These data provide convincing evidence that the native toxin used in anisotropy decay should fully occupy the binding sites in the Fl-T177C conjugated AChBP at the micromolar concentrations used in those experiments.


Figure 3
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  		FIGURE 3.
Determination of ligand binding to Fl-T177C. Panel A, steady-state emission of fluorescein was monitored in the presence of TMR-Bgt until fluorescence energy transfer was maximal (200 nM). Total binding sites = 100 nM. Panel B, binding of TMR-Bgt as determined by fluorescence quenching indicated in panel A. KD was determined from the negative inverse slope of the linear fit of the Scatchard Plot (inset), where bound = 1 x 107 (ymax - y[x])/(ymax - ymin), where y[x] = fluorescence signal at concentration x, and free = total TMR-Bgt added minus the bound fraction. Linearity in the Scatchard plot reveals little or no difference in affinity between fluorescein-modified and unmodified sites. Panel C, reversal of TMR-Bgt quenching of fluorescein, upon the addition of 100 nM increasing concentrations of epibatidine. [Binding sites] = 200 nM. cps, counts/s.

 
To characterize further ligand binding to the Fl-T177 conjugate, epibatidine was added in incremental concentrations to compete with bound TMR-{alpha}-bungarotoxin and promote its dissociation (Fig. 3C). Using 200 nM binding sites at 500 nM epibatidine, a complete reversal of the fluorescein quenching was observed. Because the competing TMR-{alpha}-bungarotoxin, present at 700 nM, was ~15-fold over its dissociation constant, half-maximal dissociation by epibatidine competition would require its concentration to be at least 15-fold over its dissociation constant. Nearly complete saturation of epibatidine would require an additional 10-fold concentration increase over its dissociation constant. From this titration epibatidine has a dissociation constant for the Fl-labeled interface of ≤3nM (Table 2), again indicating that these ligands will saturate all sites in AChBP in anisotropy assays.

In two other combinations of mutants and ligands, Fl-D194C with epibatidine and Fl-Y164C with {alpha}-bungarotoxin, no significant change in fluorescence anisotropy was observed. As with the Fl-Cys-188 mutant, we were fortunate that in both of these cases binding of the ligand in question resulted in a large enhancement of steady-state fluorescence of the conjugated fluorescein. We again used stopped-flow measurements to determine rates of association and dissociation by monitoring changes in fluorescein emission. Dissociation constants determined in this manner were 16 nM for epibatidine binding to Fl-D194C and 14 nM for {alpha}-bungarotoxin binding to Fl-Y164C (Table 2). Hence, in anisotropy assays, the lack of observed change in backbone mobility is not explained by a lack of ligand occupation. To verify that the approximate KD values obtained in the TMR-Bgt experiments with Fl-T177C yielded equivalent results, we used the FRET method to examine the interaction of Fl-Y164C with the TMR-labeled toxin. In this experiment we are limited to a lower threshold of 100 nM binding sites, so although we were only able to determine with confidence that our KD for TMR-Bgt was ≤120 nM, these findings are consistent with the stopped-flow data for the same conjugated mutant AChBP.

Time-resolved Fluorescence Anisotropy Decay of Apo-AChBP—To map {alpha}-carbonyl backbone flexibility around the agonist/antagonist binding sites and monitor ligand-induced changes in this flexibility, the sulfhydryl-reactive fluorophore MTS-Fl was selectively conjugated to substituted cysteine residues and nondisulfide-bonded Cys-188 in separate AChBP mutants. Time-resolved fluorescence anisotropy decay of each conjugate was monitored in the absence and presence of nicotinic agonists and antagonists. This approach typically distinguishes up to three types of rotational-diffusion processes in proteins; they are very fast and irresolvable anisotropic fluctuations associated with the fluorophore undergoing torsional movement about its linker arm and amino acid side chain (<1 ns), fast anisotropic motions that largely correspond to local fluctuations in the {alpha}-carbon backbone around the site of conjugation ({varphi}1, ~1-10 ns), and slower, isotropic global rotational diffusion of the entire biomolecule ({varphi}2) (18, 20-24). The determination of the rotational correlation times ({varphi}1 and {varphi}2) and their amplitudes (beta1 and beta2) associated with these diffusional processes is inherently challenging particularly when the emission lifetime of the reporter group is many times faster than the rotational correlation time of the biomolecule studied as is the case here with MTS-Fl ({tau} ~ 3-4 ns) and AChBP ({varphi}r ~ 120 ns). To reduce the uncertainty in the fitting analysis results, the {varphi}2 parameter was allowed to either float or be constrained to the {varphi}2 values that had been previously determined with a longer-lifetime fluorescent probe (124 ns for apo and small ligand-bound AChBP or 142 ns for {alpha}-bungarotoxinbound AChBP) (18). With the exception of the T177C mutant results, constraining or floating {varphi}2 yielded comparable {chi}2r values and no systematic variation of the residual plots. Reasoning that constraining {varphi}2 to the previous experimentally determined values would produce better estimates of the fast anisotropy decay parameters, only the results from the constrained fits are reported here for all the samples except the T177C mutant. Because of its complex and extensive mobility, the T177C data were best fit when all the anisotropy parameters were unconstrained.

Comparative Residue Analysis of Segmental Motion in AChBP—The results from the fitting analysis described under "Experimental Procedures" are summarized in Table 3 and Fig. 1B. The ratios of the fractional amplitude to fast rotational correlation time (fxb/{varphi}1), which generally reflect the rate and/or range of segmental motions around each site of conjugation, strongly suggest a rank order of {alpha}-carbon backbone mobility of Fl-T177C > Fl-K139C > Fl-N158C > Fl-Y164C > Fl-D194C > Fl-C188. Fl-Cys-188 in the beta9-beta10 hairpin loop (loop C) and Fl-D194C in the beta10 strand, a residue in contact with the protein core based on the crystal structures (5, 9, 25), were associated with the lowest segmental mobility of the sites examined. In contrast, Fl-T177C in the beta9 strand, which is part of a beta sandwich that is primarily stabilized by its interaction with the beta10 strand, was associated with the greatest segmental motion. The characteristics of the rotational diffusion of the reporter group at T177C are more complex than the other sites examined, because constraining its {varphi}2 to the previously determined values yielded unacceptable fits and suggests fast, large amplitude excursions of its transition moment that are more complex than with the other conjugates. Accordingly, the complexity of the diffusion and the relatively rapid depolarization of the emission from the T177C conjugate is consistent with a highly mobile segment. Sites in the beta8-beta9 loop (Fl-Y164C and Fl-N158C, loop F), a region poorly resolved in several crystal structures, displayed comparable and intermediate segmental flexibilities. Fl-K139C in the beta7 strand appeared to be associated with considerable segmental flexibility.


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  		TABLE 3
Time-resolved fluorescence anisotropy decay parameters for FI-AChBP conjugates

Multiexponential anisotropy decay profiles from fluorescein-labeled AChBP mutants were fit to Equation 3, where beta1 and beta2 are the magnitudes of the anisotropy decay associated with the fast and slow diffusion processes, respectively, {varphi}1 is the rate of anisotropy decay associated with the fast process, {psi}2r is the reduced {varkappa}2 of the fitting analyses, and <{tau}> is the amplitude-weighted average fluorescence lifetime. fxb/{varphi}1 is a complex function of the rate and range of motion associated with the fast diffusional process, where larger values correspond to increased flexibility.

 


Figure 4
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  		FIGURE 4.
Time-resolved fluorescence anisotropy decay for the fluorescein-conjugated AChBP mutants. Panel A, Fl-K139C (0.5 µM). Panel B, Fl-N158C (2 µM). Panel C, Fl-T177C (1 µM). Panel D, Fl-D194C (1.4 µM). Experiments were run either in the absence of ligand (Apo) or in the presence of epibatidine (Epi; 3.3 µM for Fl-K139C and Fl-T177C, 6.7 µM for Fl-N158C, and 15 µM for Fl-D194C), d-tubocurarine (d-Tubo;33 µM for Fl-K139C and Fl-T177C, 67 µM for Fl-N158C, and 100 µM for Fl-D194C), or {alpha}-bungarotoxin (Bgt; 3.3 µM for Fl-K139C and Fl-T177C, 6.7 µM for Fl-N158C, and 13 µM for Fl-D194C).

 
Effects of Ligand Binding on Anisotropy Decay Parameters—Initially, all the Fl conjugates were screened with one agonist, epibatidine, and two antagonists, d-tubocurarine, an alkaloid, and {alpha}-bungarotoxin, an 8-kDa peptidic {alpha}-neurotoxin (Fig. 2). Fl-K139C, located in the beta7 strand on the membrane side of the binding pocket on the principal side of the subunit interface, was stabilized by binding of the two antagonists but not significantly affected by the agonist epibatidine. This is most easily seen in the anisotropy decay profiles (Fig. 4A) and in the fxb/{varphi}1 values (Table 3) where d-tubocurarine and {alpha}-bungarotoxin decreased fxb/{varphi}1 from 0.21 to 0.15 and 0.14, respectively.

Fl-N158C, located in the apical portion of the F-loop on the complementary side of the subunit interface, was significantly stabilized by {alpha}-bungarotoxin binding but was mobilized by both epibatidine and d-tubocurarine (Fig. 4B). {alpha}-Bungarotoxin decreased fxb/{varphi}1 from 0.19 to 0.05, whereas epibatidine and d-tubocurarine increased fxb/{varphi}1 from 0.19 to 0.30 and 0.26, respectively (Table 3). The dramatic stabilization of Fl-N158C by {alpha}-neurotoxin is consistent with the crystal structure of {alpha}-cobratoxin bound to AChBP that shows loop II of the {alpha}-neurotoxin interacting with this segment of loop F (9). The lack of stabilization by the relatively large alkaloid d-tubocurarine suggests that it does not interact directly with this more apical portion of loop F. Fl-T177C, located in the beta9 strand, was highly flexible as discussed above, and its mobility was unaffected by any of the ligands assayed (Fig. 4C and Table 3). For Fl-D194C on the beta10 strand, epibatidine binding had no significant effect on the fast mobility of the reporter, whereas d-tubocurarine binding increased fxb/{varphi}1 from 0.12 to 0.21 and {alpha}-bungarotoxin decreased fxb/{varphi}1 from 0.12 to 0.05 (Fig. 4D and Table 3).

The site on loop C was of particular interest since agonists induced a closing movement of loop C toward the core of the protein (8), and this conformational change may be associated with propagation of the agonist binding event to the ion channel gate in the transmembrane domain (3). Consequently, we examined Fl-C188, positioned at the tip of loop C, with several ligands, but differences in the decay of anisotropy did not correlate with the behavior of the respective compounds as agonists and antagonists (Fig. 5A). In the absence of any ligand the value of the fxb/{varphi}1 parameter was 0.04. The addition of three agonists, epibatidine (0.06), nicotine (0.09), and 4-OH,2MeOBA (0.11), and one antagonist, MLA (0.19), increased segmental flexibility, whereas the addition of another agonist, lobeline (0.02), decreased mobility as measured by the fxb/{varphi}1 parameter (Table 3). Carbamylcholine, an agonist, and {alpha}-bungarotoxin (0.03) were without significant effect (Table 3). Accordingly, mobility parameters of loop C when modified by removal of one of the two vicinal cysteines for fluorophore conjugation were not correlated with the pharmacologic actions of the ligands.


Figure 5
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  		FIGURE 5.
Time-resolved fluorescence anisotropy decay for the fluorescein-conjugated AChBP mutants. Panel A, Fl-Cys-188 (1.2 µM). Panel B, Fl-Y164C (1.5 µM). Experiments were run either in the absence of ligand (Apo) or in the presence of epibatidine (Epi;14 µM), d-tubocurarine (d-Tubo;28 µM for Fl-Y164C and 14 µM for Fl-C188), {alpha}-bungarotoxin (Bgt;14 µM for Fl-Y164C and 36 µM for Fl-C188), carbamylcholine (Carb;28 µM for Fl-Y164C and 140 µM for Fl-C188), nicotine (Nic;14 µM), lobeline (Lob;14 µM), MLA (14 µM), or 4-OH,2MeOBA (28 µM for Fl-Y164C and 2.4 µM for Fl-C188).

 
Initial screening of Fl-Y164C, located in the COOH-terminal portion of loop F approaching the membrane, suggested a possible distinct agonist response with epibatidine, an agonist, increasing flexibility dramatically (fxb/{varphi}1 increased from 0.16 to 0.29), and the antagonist, d-tubocurarine, decreasing the flexibility (fxb/{varphi}1 decreased from 0.16 to 0.10), whereas the larger peptide antagonist, {alpha}-bungarotoxin, produced no significant effect on mobility (fxb/{varphi}1 = 0.15) (Fig. 5B, Table 3). Four additional agonists were assayed, nicotine, carbamylcholine, lobeline, and 4-OH,2MeOBA (an {alpha}7-selective partial agonist); all increased {alpha}-carbon backbone mobility in this loop F region, with their fxb/{varphi}1 values increasing from 0.16 to 0.23, 0.22, 0.21, and 0.22, respectively (Table 3). MLA, an antagonist, behaved like d-tubocurarine and decreased mobility; the fxb/{varphi}1 value decreased from 0.16 to 0.11 (Table 3), confirming that agonists increase, and antagonists either have no effect or decrease segmental mobility of the segment of loop F proximal to the membrane.

Assuming the difference between the fundamental anisotropy for fluorescein (0.35) and the sum of the observed decay amplitudes (beta1 + beta2; Table 3) represents the maximum angular excursions of the very rapid linker-arm (or tether-arm) motions, then ligands only significantly affected the maximum angular excursions of fluorescein attached to the N158C and Cys-188 AChBP mutants. For both of these conjugates {alpha}-bungarotoxin uniquely produced very significant decreases in the amplitudes of the very rapid excursions. In the case of the N158C conjugate, the very rapid decay amplitude decreased from 0.119 to 0.069, and for the Cys-188 conjugate it decreased from 0.82 to 0.053. These results are consistent with either a direct "interaction" of the bound-toxin with the reporter group or a toxin-induced conformational state that is associated with restrictions of very fast torsional motions of conjugated fluorescein. In the case of the C187S mutant, the 80-fold decrease in {alpha}-bungarotoxin affinity for the Fl-Cys-188 conjugate over the unlabeled mutant makes the case for a direct interaction more probable.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Using Lymnaea AChBP as a surrogate of the extracellular domain of the nAChR, we combined site-directed labeling with time-resolved fluorescence anisotropy to measure the effects of agonists and competitive antagonists on the {alpha}-carbon backbone flexibility at six sites near the agonist binding pocket. Control experiments demonstrated that the conjugated fluorescent probes did not block ligand occupation under the conditions of the anisotropy measurements. Specifically, we monitored anisotropy decay from fluorescein conjugated to three sites in the beta9-beta10 sandwich as well as a site in the beta7 strand on the membrane side of the binding pocket in the principal subunit face and two sites in the loop connecting the beta8 and beta9 strands across from the binding pocket in the complementary subunit face (Fig. 1). With the exception of the highly flexible site in the beta9 strand, both agonists and antagonists perturbed the backbone flexibility of each site examined; however, agonist-selective changes were only observed in the complementary subunit in a segment of the beta8-beta9 loop at Y164C (in loop F). More precisely, agonists and partial agonists increased the range and/or rate of the fast anisotropy decay processes that are usually associated with the {alpha}-carbon backbone fluctuations. This agonist-selective change in conformational dynamics probably occurs across fresh water and marine species in the animal kingdom, because we observe comparable agonist-specific differences in anisotropy decay for the homologous site (S167C) in loop F of AChBP from Aplysia californica (data not shown).

Although x-ray crystallographic studies have not revealed ligand-dependent changes in the conformation of loop F, solution-based studies have suggested structural and/or functional links between ligand binding and loop F. With AChBP, hydrogen-deuterium exchange analysis (26) and steady-state spectrofluorometric measurements using acrylodan conjugates (16) have identified ligand-specific changes in the solvent accessibility of loop F. Accessibility studies with cysteine-substituted mutant {gamma}-aminobutyric acid, type A receptors (27) and site-directed mutagenesis with the 5-HT3 receptor (28) also indicate state-dependent changes in the solvent exposure of loop F. Additionally, hydrophobic photolabeling of the nAChR loop F occurs in the open state of the receptor but not during the closed or desensitized states (29), indicating that this region likely moves during channel gating. Also, labeling of the portion of loop F proximal to the membrane occurs only when agonist is bound (30), again indicating a potential agonist-specific movement in a portion of loop F some distance from the binding pocket. Furthermore, mutation of a single residue in the beta8-beta9 loop of the mammalian {alpha}4beta2 nAChR ({alpha}E180Q), homologous to Arg-170 in AChBP) abolishes Ca2+ potentiation of the ACh response (31). Taken together the above observations show loop F to be a dynamic entity that most likely adopts conformations not evident from crystallographic structures. Our data with the Fl-Y164C conjugate support the notion that an agonist activation signal is transmitted to a portion of loop F. Within the limitations of our soluble surrogate protein as a model, we can only demonstrate that loop F responds with a change in conformational dynamics when agonists bind at their recognition site. Establishing the involvement of loop F in the direct transmission of a signal to the ion gate requires confirmation with intact receptors.

The observed, localized agonist-selective increase in backbone flexibility surrounding Fl-Y164C is most likely propagated across the principal-complementary subunit interface rather than within the beta9-beta10 sandwich from the principal-subunit beta9-beta10 hairpin (loop C). The reasons for this conclusion are, first, that no agonist-selective changes in the anisotropy decay rates of the reporter groups on either the beta9-beta10 hairpin or the beta9 strand were observed. Indeed, the segment around T177C in the beta9 strand is so flexible that no ligand examined produced measurable changes in the anisotropy decay, and it seems unlikely that conformational signals can be carried by highly dynamic, unconstrained structural elements. Additionally, the relative proximity of Y164C to the binding pocket across the nearby subunit interface compared with the relative long distance to loop C in the principal subunit make trans-subunit signal propagation a more reasonable alternative.

How agonist binding increases the rate and/or range of backbone motions of part of the beta8-beta9 loop is unclear. One way to enhance flexibility of a surface element would be through a reduction in the element's interaction with neighboring structural elements. The nearest structural elements to the beta8-beta9 loop are an antiparallel beta sheet and the beta9-beta10 hairpin. The antiparallel beta sheet is formed by beta5, beta6, beta2, and beta1 strands with the beta1 strand running almost parallel to most of the beta8-beta9 loop. Overlaying the x-ray structures of nicotine- and bufferbound crystal structures (PDB access codes 1I9B [PDB] and 1UW6) reveals little or no agonist (nicotine) specific perturbation of this beta sheet, suggesting that conformational changes in the beta sheet probably do not cause changes in the flexibility of loop F. Additionally, although the beta9-beta10 hairpin moves inward toward the protein core with agonist binding, there does not appear to be any hydrogen or other non-covalent bonding with any part of the beta8-beta9 loop with or without nicotine. That said, examination of the B factors shows nicotine binding is associated with lower B factors in the beta9-beta10 hairpin upon binding and, interestingly, increased B factors and less secondary structure along most of the beta8-beta9 loop. Although this is largely consistent with the present results, it does not explain the physical basis of the nicotine-elicited differences in the beta8-beta9 loop conformation.

In addition to elucidating ligand-induced changes in conformational dynamics, the results provide insight into the solution backbone dynamics of AChBP that is unobtainable from crystallographic B factors. B factors provide a measure of dispersion of each atom at rest without any time reference and, therefore, do not resolve static from dynamic disorder let alone reveal the time domain in which atomic fluctuations occur. Site-directed labeling combined with time-resolved fluorescence anisotropy, although modifying the native protein, on the other hand, provides information on the solution dynamics of reporter groups attached to areas of interest in the picosecond-nanosecond time domain. Here, of course, one assumes that the conjugated cysteine-substitution mutant displays largely the same conformational dynamics as the wild-type molecule. With this caveat in mind, the present results reveal the beta9-beta10 sandwich to be dynamically heterogeneous with the beta10 strand relatively anchored and undergoing limited and slow fluctuations in the picosecond-nanosecond time domain, whereas the beta9 strand is highly dynamic in this time domain. This is a reasonable conclusion because the beta10 strand is extensively hydrogen-bonded to the protein core, whereas the beta9 strand is not. Furthermore, assuming that the beta9-beta10 sandwich remains intact in the solution state, the apparent dynamic character of the beta9 strand (as revealed in the anisotropy decay of Fl-T177C) probably reflects the unique characteristics of the motion of a reporter group attached to one strand of a beta sandwich. These strands are primarily linked to the protein core through antiparallel hydrogen bonds to its partner (beta10) that is extensively bonded (non-covalently) to the protein core. In such a situation the minimally constrained beta strand (beta9) should be able to undergo fast butterfly like motions that allow very large and rapid angular excursions of the reporter group producing rapid, large-amplitude depolarization.

Although our data are not consistent with the activation signal being propagated down the beta9-beta10 sandwich, the beta9-beta10 hairpin (loop C) plays a critical role in agonist binding. Similar to prior findings on the nAChR, we observe a large loss in binding affinity for both epibatidine and {alpha}-bungarotoxin upon reduction or removal of the loop C vicinal disulfide. Nevertheless, even with the loss in affinity, reduction and labeling of the vicinal cysteines resulted in functional channels (32, 33). Thus, an intact loop C structure appears to play an important role in ligand binding energetics but may still allow appreciable activation when disrupted. That other Cys-loop family members do not have vicinal cysteines in the loop C tip but likely have a conserved activation mechanism supports the notion that loop C may provide binding affinity for cholinergic ligands but not be strictly essential for channel gating. Hence, the anisotropy decay results from Fl-Cys-188 may be explained by thinking of stabilization that comes from binding of a given ligand as being due to an affinity of that ligand for loop C. Lobeline, for example, stabilizes loop C, consistent with the crystal structure showing loop C packed down tightly around the ligand (8). MLA, in contrast, results in the greatest increase in loop C mobility; the crystal structure of this complex displays loop C in varying degrees of closure around the binding pocket, which supports limited interaction of the ligand with the loop. Stabilization by MLA likely occurs elsewhere in the binding site. Epibatidine is intermediate in its effects on flexibility at this site, indicating it derives moderate affinity through interactions with loop C.

Our results with the Fl-Cys-188 conjugate should be viewed cautiously. First, 1H,15N heteronuclear single quantum correlation spectroscopy of [15N]cysteine-substituted AChBP indicate conformational heterogeneity of loop C cysteines (12). Time-resolved fluorescence anisotropy reveals a cumulative signal from all five subunits. Second, the elimination of the loop C disulfide bond with the C187S mutation undoubtedly changes the structure of loop C. Finally, the Fl-Cys-188 fast anisotropy decay parameters indicated less backbone mobility than the reporter groups in the beta9 (Fl-T177C) and beta10 (Fl-D194C) sheets that form loop C (beta9-beta10 hairpin); this low segmental mobility may result in part from the conjugated fluorescein forming a surface interaction between loop C and the core of the protein. Our attempts to label engineered cysteines at positions in loop C that would not require breaking the vicinal disulfide resulted in lack of expression, likely due to misfolding and possibly a disulfide rearrangement.

With regard to the other sites examined, the anisotropy decays associated with the Fl-N158C and Fl-Y164C conjugates show the beta8-beta9 loop to be in a significantly more dynamic state than the constrained beta10 strand (as reflected in the anisotropy decay of Fl-D194C). Similarly, the beta7 strand in the absence of any ligand appears to be flexible and exhibits greater amplitudes for the fast phase of decay. The significance of the mobility of these sites is unclear, but flexible binding surfaces can increase the rate of ligand binding, broaden substrate specificity, and enhance free energy of binding by optimizing non-covalent interactions between protein and ligand, thereby increasing enthalpy. Moreover, binding-induced increases in flexibility may minimize the entropy loss associated with binding and, thus, also increase the free energy of binding.

Various allosteric mechanisms and structural linkages have been proposed for agonist activation of the channel gate. Structural work from electron micrographs of the full-length nAChR led to the hypothesis of an agonist-induced 10-15° rotation of the core of the extracellular beta-sandwich of only the principal subunits about an axis normal to the plasma membrane (11, 34). More recently, Auerbach and co-workers (35-37) tested this hypothesis with a combination of electrophysiology and mutagenesis and found no evidence to support a synchronous rigid body movement in the extracellular domain of the receptor. This mutagenesis work was consistent with agonist-induced movements in both the beta1-beta2 linker and the beta6-beta7 linker (known as the Cys loop), proximal to the membrane interface, after a movement in the beta4-beta5 linker that constitutes a portion of the agonist binding site (Tyr-89). Our time-resolved fluorescence results reveal regional variations in mobility that are consistent with this hypothetical sequential mechanism of activation but do not offer evidence for the synchronous rigid body rotation hypothesis. Because the anisotropy decay does not measure events slower than global rotation of the pentameric protein, we cannot exclude ligand-elicited rigid body motions. Nevertheless, the changes in conformation reflected in segmental motion distal to the agonist binding site reveal segmental mobility and communication across the subunit interfaces as being potentially critical for the activation process.

In addition to generating evidence for a functional role of loop F conformational dynamics in ligand activation and a potential trans-subunit propagation of the agonist binding signal across the principal/complementary subunit interface, our results show AChBP and presumably the nAChR extracellular domain to be a dynamic entity. AChBP exhibits a wide range of segmental fluctuations between discrete regions in the {alpha}-carbon backbone. Structurally and pharmacologically distinctive ligands exert disparate effects on the stability of different domains of AChBP well removed from the binding site. This information on backbone flexibility expands our understanding of the solution behavior of the existing model of nAChR extracellular domain structure and should be applicable in drug design. The crystal structures provide a critical starting template for study; decay of fluorescence anisotropy enables one to examine the dynamic dimensions of structure in solution within the picosecond-nanosecond time frame. Additionally, the apparent principal-complementary transmission of the activation signal to part of the beta8-beta9 loop (loop F) is consistent with concerted models of nAChR activation, which require intersubunit communication.


   FOOTNOTES
 
* This work was supported by National Institutes of Health Grant R37-GM18360 (to P. T.) and a pre-doctoral fellowship from the Pharmaceutical Research Manufacturers Association Foundation (to R. E. H.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement"in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

1 To whom correspondence should be addressed: Division of Biomedical Sciences, B605 Statistics Rd., University of California Riverside, Riverside, CA 92521-0121. Tel.: 951-827-3831; Fax: 951-827-5504; E-mail: david.johnson{at}ucr.edu.

2 The abbreviations used are: nAChR, nicotinic acetylcholine receptor; AChBP, acetylcholine-binding protein; MTS-Fl, 2-[(5-fluoresceinyl)aminocarbonyl]ethyl methanethiosulfonate; TMR-Bgt, tetramethylrhodamine-{alpha}-bungarotoxin; MLA, methyllycaconitine; 4-OH,2MeOBA, 4-hydroxy, 2-methoxy-benzylidene anabaseine; FRET, fluorescence resonance energy transfer; WT, wild type; 5-HT3, 5-hydroxytryptamine 3. Back



   REFERENCES
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 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
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