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NMR-based Binding Screen and Structural Analysis of the Complex Formed between α-Cobratoxin and an 18-Mer Cognate Peptide Derived from the α1 Subunit of the Nicotinic Acetylcholine Receptor fromTorpedo californica *
To whom correspondence should be addressed: Dept. of Molecular Pharmacology, Physiology, and Biotechnology, Brown Medical School, Box G-B391, Providence, RI 02912. Tel.: 401-863-1034; Fax: 401-863-1595
* This work was supported by Research Grants GM32629 and NS34348 (to E. H.) from the National Institutes of Health.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.The atomic coordinates and the structure factors (code 1LXG and 1LXH ) have been deposited in the Protein Data Bank, Research Collaboratory for Structural Bioinformatics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org/). ‡ Performed this work in partial fulfillment of the requirements for a Ph.D. from Brown University.
The α18-mer peptide, spanning residues 181–198 of the Torpedo nicotinic acetylcholine receptor α1 subunit, contains key binding determinants for agonists and competitive antagonists. To investigate whether the α18-mer can bind other α-neurotoxins besides α-bungarotoxin, we designed a two-dimensional1H-15N heteronuclear single quantum correlation experiment to screen four related neurotoxins for their binding ability to the peptide. Of the four toxins tested (erabutoxin a, erabutoxin b, LSIII, and α-cobratoxin), only α-cobratoxin binds the α18-mer to form a 1:1 complex. The NMR solution structure of the α-cobratoxin·α18-mer complex was determined with a backbone root mean square deviation of 1.46 Å. In the structure, α-cobratoxin contacts the α18-mer at the tips of loop I and II and through C-terminal cationic residues. The contact zone derived from the intermolecular nuclear Overhauser effects is in agreement with recent biochemical data. Furthermore, the structural models support the involvement of cation-π interactions in stabilizing the complex. In addition, the binding screen results suggest that C-terminal cationic residues of α-bungarotoxin and α-cobratoxin contribute significantly to binding of the α18-mer. Finally, we present a structural model for nicotinic acetylcholine receptor-α-cobratoxin interaction by superimposing the α-cobratoxin·α18-mer complex onto the crystal structure of the acetylcholine-binding protein (Protein Data Bank code 1I9B).
nicotinic acetylcholine receptor
heteronuclear single quantum correlation spectroscopy
total correlation spectroscopy
nuclear Overhauser enhancement spectroscopy
nuclear Overhauser effect
root mean square deviation
Crystallography and NMR System
The nicotinic acetylcholine receptor (nAChR)1 mediates excitatory transmission at the neuromuscular junction and in the central and peripheral nervous systems. As the prototype for the superfamily of ligand-gated ion channels, it has been intensively studied. The ligand-gated ion channel family includes the glycine, γ-aminobutyric acid, type A, γ-aminobutyric acid, type C, and 5-hydroxytryptamine type 3 receptors. All are apparently pentamers composed of either identical or homologous transmembrane subunits. The skeletal muscle-type nAChR contains two α1 subunits and one each of the β1, γ(ε), and δ subunits. The αγ(ε) and αδ interfaces form the sites for ligand binding (see Ref.
for review). Recently, a natural homologue of the extracellular domain of the nAChR, the acetylcholine-binding protein (AChBP), was discovered as a soluble, secreted pentamer in the central nervous system of the snail, Lymnaea stagnalis. The x-ray crystal structure of this AChBP at 2.7 Å reveals the putative conformational architecture of the extracellular domain of the nAChR (
The snake venom-derived α-neurotoxins are classic competitive antagonists of the muscle-type nAChRs, and they can be divided into two related groups as follows: short and long α-neurotoxins. X-ray crystal structures and NMR solution structures of several different α-neurotoxins reveal a conserved structural pattern consisting of a core of disulfides with three fingers or loops (
) have helped elucidate the interactions between the nAChR and the α-neurotoxins and have led to new models based on the docking of a rigid structural model of an α-neurotoxin to the proposal ligand-binding site (
α-Cobratoxin (Cbtx), a member of the long α-neurotoxin family, is obtained from the venom of Naja naja kaouthia (previously called Naja naja siamensis). Cbtx has 71 amino acid residues and 5 disulfide bridges.
It binds muscle-type nAChR at the neuromuscular junction and causes paralysis by preventing acetylcholine binding to the nAChR. LSIII, from the venom of Laticauda semifasciata, is also classified as a long α-neurotoxin because of its fifth disulfide bond and is composed of 66 amino acid residues (
) indicates that residues 173–204 from the α1 subunit of the nAChR form a major binding determinant for α-bungarotoxin (Bgtx), a long α-neurotoxin from Bungarus multicinctus. A peptide fragment (α18-mer) with a sequence corresponding to residues 181–198 (α1-181YRGWKHWVYYTCCPDTPY198), from the Torpedo californica nAChR, binds Bgtx with an apparentKd of ∼65 nm (
). The structures of these Bgtx·peptide complexes provide valuable information about the contribution of the contact residues involving in Bgtx binding to the native nAChR. We now report an NMR-based binding assay using the α18-mer to screen for interaction with four α-neurotoxins related to Bgtx. We also report an NMR structural analysis of the Cbtx·α18-mer complex. Our goal is to further define the contribution of individual amino acid residues in the α1 subunit to the interaction between the nAChR and the α-neurotoxins.
Expression of the α18-Mer Peptide
The α18-mer was expressed and purified as described previously (
). Briefly, the peptide was expressed as a ketosteroid isomerase fusion protein using plasmid pET 31b(+) in standard M9 medium and Escherichia coli strain BL21 (DE3) (Novagen) with the replacement of15NH4Cl for NH4Cl to uniformly enrich the peptide with 15N. The peptide was cleaved from the fusion protein by reaction with cyanogen bromide, and it was then purified by reverse phase-high performance liquid chromatography. As before, the C-terminal homoserine lactone form of the α18-mer was used for the NMR studies described below.
NMR Sample Preparation
For the HSQC screen, the15N-enriched α18-mer was resuspended in 50 mmof per-deuterated potassium acetate buffer (pH 4.0) with 5% D2O and 0.05% sodium azide. Cbtx, Ea, Eb, and LSIII (from Sigma) were prepared in the same buffer at a concentration of 1 mm. Toxins from their stock solutions were added to the α18-mer in a 1:1 stoichiometry. The final concentration of all mixtures was 0.2 mm. For the structure analysis of the Cbtx·α18-mer complex, the 15N labeled α18-mer was resuspended in the buffer described above. Cbtx, from a 5 mm stock solution, was then added to the α18-mer to form a 1:1 Cbtx·α18-mer complex. The final concentration of the Cbtx·α18-mer complex was 1.6 mm.
All NMR spectra were recorded on a Bruker Avance 600 MHz NMR spectrometer at a temperature of 35 °C. Chemical shifts at this temperature were calibrated with respect to internal 3-(trimethylsilyl) tetradeutero sodium propionate (0.0 ppm). The various toxin/α18-mer samples were analyzed by a two-dimensional15N heteronuclear single quantum correlation (1H-15N HSQC) experiment (
). The formation of the Cbtx·α18-mer complex was confirmed in a mole ratio titration series using the HSQC protocol for analysis. Amino acid spin systems were identified by two-dimensional total correlation spectroscopy (TOCSY) (
) using a 60-ms MLEV-17 spin-lock sequence. The assignments of the HN protons and Hα protons of the amino acid spin systems of the peptide were further confirmed by a three-dimensional HNHA experiment (
) with a mixing time of 120 ms. NH exchange experiments in D2O were performed to identify slowly exchanging amide protons presumably involved in stable hydrogen bonds. The Cbtx·α18-mer complex sample was lyophilized, and immediately after resuspending the complex in 99.99% D2O, an HSQC and 12 sequential TOCSY spectra (3-h acquisition time for each) were collected. All NMR spectra were processed and analyzed with XwinNmr (Bruker), NMRPipe (
The distance constraints were obtained from the cross-peak volumes in the two-dimensional NOESY spectra of the Cbtx·α18-mer complex by integration using the Gaussian fitting protocol in Sparky. The cross-peaks, according to their volumes, were manually classified into three categories: strong, medium, and weak with corresponding distance ranges of 1.8–3.0, 1.8–4.0, and 1.8–5.0 Å, respectively. Those amide protons, involved in β-sheet-like Hα-Hα and Hα-HNNOEs, which could be identified in NH exchange experiments as stable to exchange after 21 h in D2O were classified as “hydrogen bond” protected amide protons and as hydrogen bond donors. For the structural incorporation of hydrogen bonds, the distance constraint of HN-O was assigned a value of 1.6–2.5 Å and that of N-O was constrained to 2.5–3.3 Å. The Hα-HN3J coupling constants of the α18-mer peptide were derived from the three-dimensional HNHA experiments (
). For 3J< 6 Hz, the dihedral angle restraint (φ) was assigned to −60 ± 30°; for 3J > 8 Hz, the φ was −120 ± 40°. All structures were calculated from random models with distance geometry and by applying the simulated annealing protocols in the Crystallography and NMR System (CNS) software package (
). The potential energy function used in these calculations was the sum of the van der Waals repulsion term whose force constant varied from 0.003 to 4 kcal mol−1 Å−4 during the cooling stage, the NOE distance constraints using a square-well potential with a force constant of 50 kcal mol−1, the dihedral angles with a force constant of 200 kcal mol−1rad−2, the bond length, and the bond angles. Pseudoatoms were used for protons that could not be stereospecifically assigned. The pseudoatom correction feature of CNS was used to adjust the NOE distance constraint range automatically. Each round of calculation was initiated with a random seed number. Resulting structural models with no NOE violation larger than 0.5 Å were regarded as acceptable structures. The 10 lowest energy structures of 100 acceptable structures were selected to form an ensemble to represent the final structural model of the Cbtx·α18-mer complex. The intermolecular contact surface areas of all the final individual Cbtx·α18-mer complex structures were calculated using Contacts of Structural Units software (
). All structure coordinates of the Cbtx·α18-mer complex have been deposited at the Research Collaboratory for Structural Bioinformatics Protein Data Bank. The entries for the ensemble structures and the minimized average structure are 1LXG and 1LXH, respectively.
), the α18-mer can form a stoichiometric complex with Bgtx that is amenable for NMR structure analysis. To investigate whether the α18-mer can also be recognized by other α-neurotoxins, we designed an NMR-based screen to test for interactions with α-neurotoxins related to Bgtx (Cbtx, Ea, Eb, and LSIII). The screen was based on the use of an15N-enriched α18-mer and a two-dimensional1H-15N HSQC experiment that is designed to acquire signal only from 15N-attached protons of the peptide (i.e. amide protons and the side chain protons of Arg and Trp). Comparison of the spectra of the α18-mer·α-neurotoxin samples with that of free α18-mer was used to determine whether there is an interaction between the α18-mer and these toxins. A binding interaction between α18-mer and the toxin is indicated in those cases where HN resonances of the α18-mer undergo chemical shift changes. An HSQC titration is necessary to determine whether this binding is in the realm of slow, medium, or fast exchange. If none of the HN resonances of the α18-mer undergo any chemical shift changes upon toxin addition, an interaction in the millimolar to sub-millimolar affinity range (the NMR sample concentration range) can be effectively ruled out.
The results shown in Fig. 1 indicate that Ea (Fig. 1A), Eb (Fig. 1B), and LSIII (Fig.1C) do not bind to the α18-mer, whereas Cbtx does form an apparent stoichiometric complex with the α18-mer (Fig.1D). We next carried out HSQC-based mole ratio titration studies to determine whether the Cbtx·α18-mer complex was indeed stoichiometric and whether the interaction is in slow exchange (Fig.2). From the HSQC titration, it is clear that the free peptide (Fig. 2A) is largely unstructured. Upon binding to Cbtx, the chemical shift positions of the HN resonances for the bound peptide are altered due to complex formation but remain fixed in position and do not vary as a function of the Cbtx concentration (Fig. 2, B–D). These observations indicate that the α18-mer forms a 1:1 complex with Cbtx, that the α18-mer acquires structure upon binding, and that the Cbtx·α18-mer complex is in slow exchange (i.e. time scale in the millisecond to second range).
Three-dimensional TOCSY-HSQC, threedimensional NOESY-HSQC, and three-dimensional HNHA experiments were performed to assign the resonances of the α18-mer in the Cbtx·α18-mer complex. From these three-dimensional NMR experiments, we assigned the observable resonances for all of the amino acid residues in the α18-mer except for the N-terminal Tyr181, which has an exchangeable HN, and the two prolines that lack amide protons. These assignments of the peptide resonances were used in the analysis of the two-dimensional NMR data to distinguish peptide resonances from Cbtx-associated proton resonances, greatly facilitating the assignment of the Cbtx resonances. Guided by the two-dimensional NMR assignments of free Cbtx (
), we completed the two-dimensional NMR assignments for free Cbtx and for the Cbtx·α18-mer complex with additional homonuclear two-dimensional NOESY and two-dimensional TOCSY experiments. By comparing the chemical shift assignments for free and bound Cbtx, we found that the region from Asp27 to Val37 of Cbtx is characterized by significant chemical shift changes upon complex formation (TableI).
Table IChemical shift perturbations in Cbtx induced by α18-mer binding
All of the resonances listed here are characterized by chemical shift changes upon peptide binding of greater than 0.15 ppm. Proton designations follow IUPAC recommended nomenclature.
Distance constraints were obtained from the intensities of the NOE peaks in two-dimensional NOESY spectra of the complex as described under “Experimental Procedures,” whereas hydrogen bonds important for secondary structure were identified by NH exchange experiments. In addition, the dihedral angle restraints for the peptide were calculated from Hα-HN3J couplings obtained through HNHA experiments. All of these constraints (TableII) were then incorporated into CNS (
) to calculate structural models of the Cbtx·α18-mer complex. The CNS calculations incorporate both distance geometry and simulated annealing protocols. After multiple rounds of calculation using different random seed numbers, a pool of 100 structures with no NOE violation larger than 0.5 Å was obtained. From the pool of acceptable structures, 10 structures with the lowest potential energy were selected to represent a structural ensemble of the Cbtx·α18-mer complex. The overall backbone atomic root mean square deviation (r.m.s.d.) between the individual structures and the mean structure of the Cbtx·α18-mer complex is 1.69 Å. We identified a highly defined region composed of residues 2–70 of Cbtx and residues 187–196 of the α18-mer. The remaining residues (residues 1 and 71 of Cbtx, 181–186 and 197–198 of the α18-mer) lacked long range NOE constraints. In this more highly defined region, the backbone r.m.s.d. is 1.46 Å (Table II). The structural ensemble of the complex is shown in Fig.3. For clarity, only the more highly defined regions are shown. The α18-mer-bound Cbtx is oriented to show the concave surface, loop I on the left, loop III on the right, and the tip of the loop II at the bottom. The overall three-finger-like motif is retained in the structure of bound Cbtx with the central β-sheet between loop II (Val19–Trp25) and loop III (Val52–Ser58) giving rise to long range Hα-Hα and Hα-HNNOEs and 6 slowly exchanging amide protons (HNs of Cys20, Thr22, Thr24, Asp53, Gln55, and Cys57). All 10 of the ensemble structures of the Cbtx·α18-mer complex contain this β-sheet. In free Cbtx, this central β-sheet is a triple-stranded one with two strands within loop II and the third strand contributed by loop III (
). However, those NOEs and accompanying slow-exchanging amide protons were not seen in the spectra of the Cbtx·α18-mer complex. We think that the binding induced conformational changes involving the tip of loop II weaken this intra-loop β-sheet to a considerable extent, so the typical evidence for a β-sheet was not observed within loop II of the peptide-bound Cbtx. Many of the large changes (>0.15 ppm) in chemical shift in bound Cbtx (Table I) indicate structural alterations in the tip of loop II (Asp27–Val37) consistent with extensive contacts between the Ile32-Arg36region of Cbtx and the peptide (Fig. 3).
Table IIStructural statistics for the Cbtx·α18-mer complex
A total of 20 intermolecular NOEs define the contact zone between the α18-mer and Cbtx (Table III). The contact zone has a surface area of ∼720 Å2 and involves the following residues: Ile9 and Thr10 in loop I of Cbtx; Ile32-Arg36 in loop II of Cbtx; Arg68 and Lys69 in the C-terminal tail of Cbtx; and Tyr189-Thr196 in the α18-mer. Tyr189 alone makes multiple contacts with Cbtx residues Ile9, Ile32, Arg33, and Lys35 (Table III). The aromatic ring of Tyr190interacts extensively with cationic residues Arg36, Arg68, and Lys69 (Table III). Intermolecular NOEs between Hα and HN of Thr196and Hα of Ile9, between the amide proton of Tyr189 and γ protons of Ile9, together with NOEs involving the side chain of Tyr189 and Hα of Ile9 indicate the formation of a hairpin-like structure in the α18-mer upon complexing with Cbtx (Table III). The hairpin-like folding is further evidenced by long range intramolecular NOEs between Trp187 and Thr196(Trp187Hα–Thr196Hγ), and between Val188 and Thr196(Val188Hα–Thr196Hγ, Val188Hβ–Thr196Hγ, and Val188Hγ–Thr196HN). Intermolecular NOEs between Thr191 and Thr10(Table III) imply that these two residues also participate in the contact between the α18-mer and Cbtx.
The proximity of aromatic residues in the α18-mer (Tyr189 and Tyr190) to several cationic residues in Cbtx (Arg33, Lys35, Arg36, Arg68, and Lys69) suggests that cation-π interactions may play an important role in the stability of the Cbtx·α18-mer complex. Following cation-π interaction analysis of the 10 structural models of the Cbtx·α18-mer complex using CaPTURE (
), we observed that 6 of the 10 structures contain energetically significant intermolecular cation-π interactions as defined by the analysis program. The following cation-π pairs were observed in the six structures: Arg33/Trp187, Arg33/Tyr189, Arg36/Tyr189, Arg36/Tyr190, Arg68/Tyr190, and Lys69/Trp187.
The backbone r.m.s.d. between the contact residues in the α18-mer (Tyr189–Thr196) in the complex and the corresponding region from the AChBP (Thr184–Ala191) is 1.37 Å. Because of this close fit, we were able to superimpose the structure of the Cbtx·α18-mer complex onto residues 184–191 in the structure of the AChBP. No additional structural manipulations were needed to obtain a structural model for the predicted AChBP·Cbtx (Fig.4) containing no steric clashes. In this initial model, statically docked loop III of Cbtx is in closer proximity to the α subunit than to the adjoining subunit, and the concave face of Cbtx is closer to the α subunit than the convex face is to the adjoining subunit. This model implies that the bulk of Cbtx is oriented, with respect to the height of the AChBP, such that it is closer to the C-terminal end than the N-terminal end of the AChBP and that it is nearly perpendicular to the long central axis of the AChBP.
By having previously documented that the α18-mer can bind Bgtx with high affinity, we wanted to test other related α-neurotoxins for α18-mer binding. We developed an NMR-based method to test for such binding. We found that the α18-mer can bind and form a stable complex with the long α-neurotoxin, Cbtx. From the observed intermolecular NOEs and the structural model of the Cbtx·α18-mer complex, we obtained important information on the points of interaction between the α18-mer and Cbtx.
Our NMR-based screen for α-neurotoxin binding to the α18-mer showed that α-neurotoxins Ea, Eb, and LSIII do not appear to interact significantly with the α18-mer. An alignment of sequences from the C-terminal tail region (Fig. 5) indicates that Ea, Eb, and LSIII all have relatively shorter C-terminal tail segments than either Cbtx or Bgtx. Importantly, they also lack cationic residues that Cbtx and Bgtx both contain in that region. Previous studies have suggested the importance of the C-terminal tail regions of long α-neurotoxins in peptide binding (
). It is likely therefore that the reason that Ea, Eb, and LSIII fail to bind to the α18-mer is due to these differences in the C-terminal tail region. It should be noted that although LSIII is categorized as a long α-neurotoxin because of the fifth disulfide bond in loop II, its C-terminal tail region, beyond the final Cys residues, is only two amino acids longer than the short α-neurotoxins, Ea and Eb. It has been reported previously that short α-neurotoxins appear, in general, to have faster on-rates (and faster off-rates) than Bgtx (
). This suggests some fundamental differences in the mode of binding among α-neurotoxins. Indeed, the results from a footprinting protection study argue that the short α-neurotoxins and Bgtx bind to opposite faces of loop C in the 187–190 region (
The interaction between Cbtx and the α18-mer is consistent with an earlier report that Cbtx can bind to a 32-mer peptide (corresponding to α1 subunit residues 173–204) with affinity similar to that of Bgtx (
). The HSQC titration (Fig. 2) results clearly demonstrate that the α18-mer can bind and form a stable 1:1 complex with Cbtx. This titration further indicated that the Cbtx·α18-mer complex is in slow exchange and that the peptide undergoes a binding induced conformational stabilization. The results obtained with the NMR-based binding screen were consistent with competition-based binding studies using 125I-Bgtx and surface-immobilized α18-mer where significant binding was evident (data not shown).
We conclude that the tip of loop I, together with loop II, and the C-terminal tail region of Cbtx form the peptide-binding pocket. This binding profile is consistent with all available NMR solution and x-ray crystal structures of complexes between Bgtx and nAChR-derived peptides (
According to the intermolecular NOEs (Table III), Ile9 and Thr10 from loop I of Cbtx are involved in α18-mer binding. This is in agreement with the structures of the Bgtx·α18-mer and Bgtx·α19-mer complexes in which, Thr8 and Pro10 (in the Bgtx·α18-mer complex) and Ala7, Ser9, and Ile11(in the Bgtx·α19-mer complex) have contacts with their respective bound peptides (
). This may imply that the tip of loop I does not form a major binding determinant for the native receptor or that one or the other of Ile9 and Thr10 is sufficient for peptide and receptor binding. The P193A mutation on α7 nAChR (Pro193 in α7 corresponds to Thr196 in the α18-mer which is in contact with Ile9) reduces the affinity for Cbtx by only 2-fold (
), suggesting that the Ala substitution mutation at this site is not sufficient to disrupt receptor recognition.
As indicated by the network of contacts between Tyr189–Tyr190 and the Ile32–Arg36 region in Cbtx, the loop II region of Cbtx forms the main contact site for the α18-mer. This is consistent with the structures of other Bgtx·peptide complexes; in those structures, Tyr189 and Tyr190 (or correspondingly, Phe186 and Tyr187, or Tyr3 and Tyr4) have extensive interactions with Leu38–Val40 of Bgtx, corresponding to Lys35–Val37 of Cbtx (
). This binding profile is also consistent with a recent double-mutant cycle analysis using Cbtx and the α7 nAChR. In this study, the R33E mutation reduced the binding affinity by 339-fold; the K35E mutation caused a 144-fold reduction in binding affinity, and the R36E mutation gave rise to a 456-fold drop in affinity (
). An earlier mutagenesis study with Cbtx indicated that the R33E mutation decreases binding affinity for Torpedo nAChR by 767-fold, whereas the R36A mutation results in a 7.4-fold reduction in affinity (
). The functional importance of Arg33 has also been pointed out in mutagenesis studies of Bgtx. The R36A mutation in Bgtx (corresponding to Arg33 in Cbtx) caused a 90-fold decrease in Bgtx binding to the mouse nAChR (
). On the receptor side, the F186E mutation in α7 (corresponding to Tyr189 in α18-mer) causes a 97-fold reduction in affinity, and the Y187H mutation (equivalent to Tyr190 in α18-mer) decreases affinity by 201-fold (
). In addition, chimeric analysis and toxin footprinting studies of nAChR α-subunits also support the involvement of Tyr189 in Bgtx binding. A Bgtx-insensitive α3 subunit can be engineered to acquire sub-micromolar affinity for Bgtx with a single K189Y mutation (
The intermolecular NOEs between Tyr190 and Arg68 and Lys69 highlight the involvement of the C-terminal tail region of Cbtx in peptide binding. This finding is consistent with the Bgtx·α18-mer structure (
), where the C-terminal tail region participates in binding. The double mutant cycle analysis involving Cbtx and the α7 nAChR together with the earlier mutagenesis studies of Cbtx, assessed by binding assays with Torpedo nAChR, also support a role for the C-terminal tail region in Cbtx binding. The F65A mutation has been shown to reduce significantly Cbtx binding affinity for α7 nAChR or Torpedo nAChR by 16- and 7-fold, respectively (
We also have focused on the cation-π interactions between the positively charged side chains of Arg and Lys and the electronegativity associated with the aromatic π cloud in the side chains of Phe, Tyr, and Trp (
). Due to the close contact between aromatic residues of α18-mer and cationic residues in Cbtx, it is not surprising that we observed energetically favorable cation-π interaction pairs in 60% of the structure models of Cbtx·α18-mer complex. Interestingly, when we used the CaPTURE program (
), we found a cation-π pairing between Arg36 and Tyr4 of the mimotope. This corresponds to the Arg33/Tyr190 pair in the Cbtx·α18-mer complex. The important roles of the cationic residues in Cbtx and of the aromatic residues in the peptide have also been indicated by mutagenesis studies, as reviewed above. Furthermore, chemical modifications of Arg34, Arg37, Arg70, and Arg72 in toxin a (Ophiophagus hannah) lead to almost complete block of binding activity to nAChR (
). This also points to the importance of cationic residues in binding to the nAChR. We suggest that cation-π interactions contribute to the high affinity interaction between Cbtx and the nAChR in a manner similar to the role of cation-π interactions in the binding of acetylcholine to the nAChR (
). Additional high resolution structural information focusing on the side chains of Cbtx and Bgtx, together with further biochemical and mutagenesis studies involving the candidate cationic residues, will be required to further evaluate this mechanism.
Because AChBP is a homologue of the extracellular domain of the nAChR (
), we used its x-ray crystal structure to generate a simple, statically docked structural model for the nAChR·α-neurotoxin complex (Fig. 4). The interaction between the nAChR and Bgtx has been modeled previously by superimposing the Bgtx·α19mer or Bgtx·mimotope complexes with AChBP (
). Here we present a nAChR·Cbtx model based on the structure of the Cbtx·α18-mer complex (Fig. 4). The orientation of Cbtx is similar to the two AChBP·Bgtx models and the recent model of the AChBP·Cbtx complex that was generated by manual docking aimed at incorporating available mutagenesis data (
). Our model, based on the structural information provided by the Cbtx·α18-mer complex, provides strong indications that Cbtx is likely to orient itself at the subunit interface, perpendicular to the long central axis of the receptor. All of these recent models indicate that α-neurotoxins approach their binding site from the periphery, rather than from the vestibule.
NMR instrumentation was funded by Grant RR08240 from the National Institutes of Health and Grant DBI-9723282 from the National Science Foundation.