We have isolated a family of insect-selective neurotoxins from the venom of the Australian funnel-web spider that appear to be good candidates for biopesticide engineering. These peptides, which we have named the Janus-faced atracotoxins (J-ACTXs), each contain 36 or 37 residues, with four disulfide bridges, and they show no homology to any sequences in the protein/DNA databases. The three-dimensional structure of one of these toxins reveals an extremely rare vicinal disulfide bridge that we demonstrate to be critical for insecticidal activity. We propose that J-ACTX comprises an ancestral protein fold that we refer to as the disulfide-directed β-hairpin.
Agricultural pesticide management is becoming increasingly complicated due to the evolution of insect resistance to classic chemical pesticides1, growing community awareness of the environmental damage caused by many agrochemicals, and strict limits enforced by many countries on the level of pesticide residuals in imported crops and livestock. These concerns have stimulated the search for ‘environmentally friendly’ pest-control strategies.
One option, although not without potential problems such as accelerated insect resistance2 and transgene escape3, is to engineer insect-specific toxins into plants. This is exemplified by the engineering of genes encoding insecticidal δ-endotoxins from the soil bacterium Bacillus thuringiensis into a variety of agriculturally important cultivars. A potentially more selective method is to use insect-specific viruses as vectors to deliver toxins to a restricted number of target species without harming beneficial insects and predators of the targeted pest. For example, most baculoviruses target only lepidopterans, an order that includes some of the most refractory agricultural pests, and this host specificity is not altered by expression of heterologous toxins4,5,6,7.
Unfortunately, there are very few well-characterized insect-specific biopesticides that lend themselves to genomic approaches. We have thus embarked on a program that aims to discover and characterize insect-specific peptide toxins suitable for engineering into plants and viruses. Here we report on a family of three homologous insecticidal neurotoxins isolated from the venom of one of the world's most lethal arachnids8, the Blue Mountains funnel-web spider Hadronyche versuta. These peptides contain 36 or 37 residues and 4 disulfide bonds, including an extremely rare vicinal disulfide — a disulfide bond between adjacent cysteine residues — that we show is essential for insecticidal activity. The toxins can be folded nonenzymatically in vitro with 100% efficiency despite the presence of four disulfide bridges, enhancing the likelihood of successful heterologous expression of the toxins in plants and virus infected insects. The three-dimensional solution structure of one of these peptides reveals an ancestral protein fold that we refer to as the disulfide-directed β-hairpin (DDH).
Isolation of toxins
A typical reverse phase (rp)HPLC fractionation of crude venom from H. versuta is shown (Fig. 1a). The venom is highly complex, and all but the earliest eluting peaks (Rt< 9 min) are peptides of mass less than 10 kDa (ref. 11). An expansion of a portion of the chromatogram (inset, Fig. 1a) shows three peptides with similar retention times. These peptides proved to be highly insecticidal (Fig. 2) but were inactive on vertebrate smooth and skeletal nerve muscle preparations (see below).
Sequencing and mass spectral analysis revealed that the three peptides are highly homologous, each is 36 or 37 residues in length with eight conserved cysteine residues involved in four disulfide bridges (Fig. 1b). We have named these peptides the J-atracotoxins (J-ACTXs); the sequences, which show no homology to any in the protein and DNA sequence databases, have been deposited in the SWISS-PROT data bank (accession numbers P82226–P82228). The LD50 values for these toxins in house crickets (167–303 pmol g−1; Fig. 2) are similar to those previously obtained for the ω-atracotoxins11.
In vitro folding of J-ACTXs
The J-ACTXs are minor components of funnel-web venom and hence sufficient amounts of native material could not be purified for detailed structural and functional characterization. Consequently, we attempted to fold synthetic J-ACTX-Hv1c (the most potent of the J-ACTXs) in vitro. Most insecticidal toxins being considered for genomic approaches to pest management have a high disulfide bridge content, such as insect-specific scorpion α-toxins4,12, which, like J-ACTX, contain four intramolecular disulfide bonds and hence 105 possible disulfide isomers. Proper oxidation/folding of these toxins in transgenic plants or virus infected insects is critical as their effectiveness could be markedly impaired if folding is inefficient. In vitro folding of these toxins is likely to give some indication of their intrinsic, sequence-dependent, nonenzymatic folding potential.
The time course of oxidation/folding of synthetic J-ACTX-Hv1c in a simple glutathione redox buffer devoid of protein folding machinery such as molecular chaperones or disulfide isomerases is shown in Fig. 3a. Remarkably, the folding reaction is almost complete within two hours and the final product, which co-elutes with native J-ACTX-Hv1c, is obtained with a 100% yield. This product was found to be as potent as native J-ACTX-Hv1c in insect bioassays, and its one-dimensional 1H NMR spectrum was identical to that of native toxin (data not shown), indicating that the native fold had been obtained. Interestingly, the folding reaction seems to occur via a discrete pathway as indicated by the three distinct peaks evident between the reduced and oxidized forms in the HPLC chromatogram, which may correspond to intermediates with one, two and three disulfides.
Three-dimensional structure of J-ACTX-Hv1c
The solution structure of the folded, synthetic J-ACTX-Hv1c was determined using standard homonuclear NMR methods13. Final structure calculations employed ∼13 restraints per residue (Table 1). An ensemble of 20 structures with the lowest residual restraint violations was used to represent the solution structure of the toxin (Fig. 4, Table 1). The structure is very precise, with backbone and heavy atom root mean square (r.m.s.) deviation values of 0.18 ± 0.06 Å and 0.57 ± 0.09 Å, respectively, for the well-defined region (residues 3–34). According to PROCHECK14, 76% of the non-Pro/Gly residues in the structured region lie in the most favored sector of the Ramachandran plot, with the remaining 24% located in the additionally allowed regions.
The structure of J-ACTX-Hv1c consists of a disulfide rich globular core comprising residues 3–19, with residues 20–34 forming a β-hairpin that projects from this region. Residues 1–2 and 35–37 at the N-terminus and C-terminus, respectively, are disordered in solution. The toxin has a high content of regular secondary structure (∼80% if the disordered N- and C-termini are excluded) as most of the polypeptide chain outside of the two β-strands (yellow in Fig. 4) is involved in various types of β-turns (green). The three central disulfide bonds form an inhibitory cystine knot (ICK) motif15,16 in which the Cys 16–Cys 32 disulfide passes through a 14-residue ring formed by the Cys 10–Cys 22 and Cys 3–Cys 17 disulfide bridges and the intervening sections of polypeptide backbone.
J-ACTX-Hv1c can be conveniently considered as comprising four loops (numbered 1–4 from the N-terminus to the C-terminus; Fig. 4c), each bounded by half-cystine residues. The hydrophobic core of the globular region is composed of the Cys 10–Cys 22 and Cys 16–Cys 32 disulfide bridges as well as the side chains of Thr 4 and Thr 20 (Fig. 4b). We previously noted that the hydrophobic core of cystine knot toxins is formed largely by the two C-terminal disulfides and a single hydrophobic residue protruding from loop 1 or 3 (Ile 5 in the case of ω-ACTX-Hv1a)17. The hydrophobic residues of J-ACTX-Hv1c are conspicuously clustered on one face of the molecule (Fig. 4b).
The vicinal disulfide bridge
The most remarkable feature of the structure of J-ACTX-Hv1c is the vicinal disulfide bridge connecting Cys 13 to Cys 14. While the disulfide configuration appeared unambiguous from the NMR data, the rarity of vicinal disulfide bonds in proteins prompted us to seek chemical confirmation. We used partial reduction at low pH followed by alkylation17,18 to trap partly reduced intermediates that would reveal the disulfide bond pattern of J-ACTX-Hv1c. In this method, partially reduced intermediates are rapidly alkylated with iodoacetamide, then the peptide is fully reduced and the remaining cysteine residues are pyridethylated. The peptides are then sequenced and the position of the disulfide bonds can be inferred from the location of the pairs of carboxamidomethylated and pyridethylated cysteines.
Two HPLC chromatograms obtained from reduction of J-ACTX-Hv1c at different temperatures are shown in Fig. 3b. Sequencing of one of the intermediates from the reaction at 37 °C yielded a peptide that was carboxamidomethylated at Cys 10, 13, 14, and 22 and pyridethylated at Cys 3, 16, 17, and 32, while one of the intermediates isolated from the reaction at 15 °C proved to be carboxamidomethylated at only Cys 10 and 22 (Fig. 4b). In combination, these intermediates provide evidence for the Cys 10–Cys 22 and Cys 13–Cys 14 disulfide bonds, thus providing chemical confirmation of the vicinal disulfide bridge.
Configuration of the vicinal disulfide bridge
Vicinal disulfide bridges in proteins are extremely rare; apart from J-ACTX, the only examples thus far reported are found in methanol dehydrogenase (MDH)19 and the α-subunit of the acetylcholine receptor (αAChR)20. Theoretical calculations21,22 predicted that the eight-membered ring formed as a result of disulfide formation between adjacent cysteines would be most stable if the intervening peptide bond was in a slightly nonplanar cis-like configuration; studies on model peptides23, as well as a low resolution structure of MDH19, appeared to confirm this view.
However, the structure of J-ACTX-Hv1c, as well as more recent high resolution structures of MDH, present a striking disparity from these theoretical and model compound analyses. A comparison of the vicinal disulfide bridge of J-ACTX-Hv1c with that from the 1.9 Å resolution crystal structure of MDH from Methylophilus W3A1 (ref. 24) is shown in Fig. 5; the atoms constituting the eight-membered ring overlay remarkably well with an r.m.s. deviation of 0.18 Å. The individual torsion angles of the disulfide ring are listed in the lower panel of Fig. 5 for J-ACTX-Hv1c, MDH24, and a disulfide bridged Cys–Cys dipeptide (cyclo-Cys-Cys)23. The Cys–Cys peptide bonds in MDH and J-ACTX-Hv1c are nonplanar (ω = −147° and −135°, respectively) but, in contrast to cyclo-Cys-Cys, they are much closer to trans than cis. The disulfide chirality is left handed for both MDH and J-ACTX-Hv1c (χ3 = −106°), again in striking contrast to cyclo-Cys-Cys where the chirality is reversed (χ3 = 94°).
The 1.94 Å resolution structure of Methylobacterium extorquens MDH25 reveals a highly nonplanar peptide bond for the vicinal disulfide bridge with a deviation of −35° from the trans configuration (that is, ω = 145°), and a recent NMR study of α-conotoxin GI revealed that the peptide bond was close to trans in a mutant with a non-native vicinal disulfide26. Thus, while only a few examples are currently available from which to formulate general conclusions, the emerging consensus appears to be that the strain imparted by formation of a vicinal disulfide bridge in native proteins is alleviated by the intervening peptide bond assuming a distorted trans configuration.
The vicinal disulfide is functionally important
The three nonvicinal disulfide bridges in J-ACTX-Hv1c connect residues that are distal in the sequence; these disulfides essentially determine the tertiary fold of cystine knot toxins27. The Cys 13–Cys 14 vicinal disulfide, on the other hand, is unlikely to play an important architectural role. Instead, the vicinal disulfide bonds in MDH and αAChR have critical functional roles: MDH activity is abolished when the vicinal disulfide at the enzyme active site is reduced19 and the vicinal disulfide bridge in αAChR is essential for acetylcholine binding28.
In order to test the functional significance of the vicinal disulfide in J-ACTX-Hv1c, we constructed a double mutant in which Cys 13 and Cys 14 were both replaced with Ser. The 1H NMR chemical shifts of the mutant toxin were almost coincident with those of native J-ACTX-Hv1c (with the exception of the mutated residues), indicating that the tertiary fold was unaffected by the point mutations. As additional proof we calculated a medium resolution backbone structure of the mutant toxin using 272 distance and 23 φ-angle restraints obtained from a single NOESY experiment. The mutant and native toxin structures overlay with a backbone r.m.s. deviation of 0.42 Å, and the overall conformation of loop 2 is largely unaffected by the two mutations in this region. However, the double mutant was completely inactive when injected into crickets at doses up to 60 times the LD50 of the native toxin (Fig. 2), demonstrating that the vicinal disulfide is critical for insecticidal activity. Thus, vicinal disulfide bridges play key functional, rather than architectural, roles in all three proteins in which they have been discovered thus far.
Is the DDH motif an ancestral protein fold?
A search of the Protein Data Bank29 revealed a number of structural homologs of J-ACTX-Hv1c, most of which are members of the ICK family of toxic polypeptides15,16. With one exception, the three best matches are with toxins that modulate voltage gated sodium channels (Fig. 6a): δ-atracotoxin27,30 from the Australian funnel-web spider; μ-agatoxin I (ref. 31) from the unrelated American funnel-web spider Agelenopsis aperta; and conotoxin GS32 from the aquatic cone snail Conus geographus.
Curiously, however, the closest structural homolog of J-ACTX-Hv1c is the cellulose binding domain of cellobiohydrolase I (CBD-CBHI) from the fungus Trichoderma reesei (Fig. 6b), which is missing the N-terminal disulfide bridge of the ICK motif. We have shown that the compact hydrophobic core of the ICK motif consists largely of the two disulfide bridges that emanate from the two β-strands that characterize the ICK fold17. The N-terminal disulfide bridge contributes very little to the hydrophobic core and, as indicated by the structure of CBD-CBHI33, is not essential to formation of the basic ICK fold. Indeed, it has been demonstrated that the tertiary structure and thermal stability of the ICK-containing trypsin inhibitor EETI II is largely unperturbed by removal of the N-terminal disulfide bridge9,34.
We propose that the ICK fold is actually a minor elaboration of a simpler ancestral fold that we refer to as the disulfide-directed β-hairpin (DDH) fold, of which CBD-CBHI is the archetypal family member. The DDH fold is shown schematically in Fig. 6c and its amino acid consensus sequence can be written as CX5–19CX2[G or P]X2CX6–19C, where X is any amino acid (Fig. 6d). The DDH fold differs from the ICK fold in that there are only two mandatory disulfide bridges (the two that form the bulk of the hydrophobic core), so that loop 1 as shown in Fig. 4c is no longer necessarily bounded by an N-terminal cysteine residue, and loop 3 is generally five residues in length with a central Gly or Pro to ensure a tight turn prior to the first β-stand. The residue following the Gly/Pro in loop 3 is generally hydrophobic. This residue along with the two buried disulfide bridges constitute the mini-hydrophobic core of this domain.
Several vertebrate proteins appear to have arisen from simple duplication of an ancestral DDH gene followed by minor loop elaborations. The ICK motif, in contrast, has not been found in vertebrates. An overlay of two molecules of J-ACTX-Hv1c on the structure of a single molecule of mamba intestinal toxin I (MIT1) from the venom of the snake Dendroaspis polylepis polylepis35 is shown in Fig. 6a. It is clear that one can obtain MIT1 by simple head-to-tail duplication of a J-ACTX-like gene followed by elaboration of loop 2 (particularly in the C-terminal DDH domain). Two molecules of J-ACTX-Hv1c can also be overlaid equally well on colipase, a molecule that is widely distributed in vertebrates and has been noted to be a structural homolog of MIT135. However, as shown in the overlay of J-ACTX-Hv1c on the C-terminal DDH domain of porcine colipase in Fig. 7b, the structural similarities can sometimes be obscured by significant loop elaborations such as the helical insertion in loop 4 of colipase.
The DDH fold differs from the previously proposed T-knot scaffold36 in that the sequence and topology are more rigidly defined for loop 3, and there is no requirement for loops 2 and 4 to have any specific orientation with respect to the β-hairpin. The DDH fold is similar to the recently proposed cystine-stabilized β-sheet (CSB) motif9. The CSB motif, however, is more rigidly defined as a triple-stranded β-sheet having the consensus sequence CX4–6CX2–6CX5–10C, which excludes many of the DDH containing proteins in Fig. 6d. Indeed, to date, no native proteins have been found that contain the elementary CSB motif without additional disulfides9.
Evolution of the DDH fold
Surprisingly, an exhaustive search of the protein and DNA sequence data bases, including numerous partially sequenced bacterial genomes, failed to identify any DDH motifs in the Archaea or Eubacteria. The DDH fold is, however, present in a wide variety of eukaryotes, including fungi, red algae (rhodophytes), arachnids, aquatic snails, plants, and vertebrates. This scenario tentatively suggests that this cystine framework evolved in an ancestral eukaryote prior to the divergence of plants, animals, and fungi, and that the marked sequence variability simply reflects the tolerance of this fold to sequence changes.
We have detected numerous ancestral duplications of this motif, as well as functional associations with other protein domains. The fungal CBD-CBHI domain is part of a multi-domain structure, and the homologous domain in the putative polysaccharide binding protein of the rhodophyte Porphyra purpurea occurs as four consecutive repeats without significant loop variability (see Pfam alignment CBD_1). Gene duplication followed by the same type of loop elaborations as observed in vertebrates can be seen in ACTX-Hvf17 from the Australian funnel-web spider10, and this duplicated motif also appears in the vertebrate embryonic head inducer Dickkopf-1 (ref. 37) where it appears to represent the N-terminal domain of a multi-domain protein10. Thus, the DDH fold appears to be widely distributed in eukaryotes, and can occur as a single domain (for example, ICK polypeptides), in duplicated form (for example, MIT1 and colipase), or as a fusion with other protein domains (for example, fungal cellobiohydrolases and insulin-like growth factor (IGF) binding proteins).
What is the physiological target of J-ACTX?
The J-ACTXs seem to be highly insect specific; at concentrations as high as 1 μM, J-ACTX-Hv1c had no effect on electrically stimulated contractions of vertebrate smooth (rat vas deferens) or skeletal (chicken biventer cervicis) muscle (data not shown), nor did it cause any adverse effects when injected into newborn mice at doses up to 3.14 μg g−1, which is five-fold higher than the LD50 in A. domesticus. However, direct application of J-ACTX-Hv1c (0.5–1.0 nmol) to the metathoracic ganglion of the cockroach Periplaneta americana caused spontaneous, uncoordinated movement of all legs within 60 s, which developed into fasciculations of the limbs and posterior sensory organs (cerci) within 1–3 min. Thus, J-ACTX-Hv1c appears to be an excitatory neurotoxin whereas, in marked contrast, ω-ACTX-Hv1a can be considered a depressant neurotoxin since it blocks reflex movements of the hind legs in this assay17.
Scorpion excitatory neurotoxins act at insect voltage gated sodium channels5. However, despite the similarity in tertiary fold between J-ACTX and the sodium channel modulators μ-agatoxin-I, δ-ACTX-Hv1, and conotoxin GS (Fig. 6a), we found that J-ACTX had no effect on whole cell potassium, sodium, or calcium currents in isolated bee brain neurons (data not shown). We also addressed the intriguing possibility that J-ACTX might mimic the α-subunit of the AChR, which also contains a vicinal disulfide bridge that forms part of the ACh binding site; AChR activity could be disrupted by J-ACTX perturbing α-subunit interactions and/or ACh binding. We found that J-ACTX-Hv1c did not affect steady state acetylcholine currents in bee brain neurons; however, it remains possible that J-ACTX antagonizes muscle AChRs or AChR subtypes present in other invertebrate neurons.
The putative bioactive surface of J-ACTX
J-ACTX possesses two strikingly dissimilar faces (Fig. 8). One face presents an almost contiguous charged surface, while the opposing face containing the vicinal disulfide is devoid of ionizable side chains (Figs 4b, 8). Thus, in the absence of a molecular target, we have tentatively named these toxins the Janus-faced atracotoxins or J-atracotoxins.
The critical functional role of the vicinal disulfide indicates that this feature is most likely present at the toxin's bioactive surface. The region surrounding the vicinal disulfide displays several features that suggest it might represent the bioactive surface of the toxin (Fig. 8). First, this region contains several exposed hydrophobic residues that are conserved in all three J-ACTXs (Ala/Thr 1, Ile 2, Ala 12, Cys 13, Cys 14, and Pro 15). Second, the side chains of Ala 12–Pro 15 form a wall that borders a hydrophobic cleft overhung by the methyl groups of Val 29. At one end of this cleft is a large cavity (Fig. 8) that might accommodate chemical groups of the target molecule.
What is the role of the vicinal disulfide bridge? One possibility is that the vicinal disulfide, while clearly not important in determining the overall fold of J-ACTX, might play a local architectural role, such as facilitating configuration of the hydrophobic cleft. However, the structure of loop 2 is largely unaltered in the Ser–Ser mutant, which argues against this hypothesis. It is possible that the redox potential of the vicinal disulfide, because of its strained cyclic geometry, might be sufficiently altered from that of typical disulfides to facilitate covalent reaction of these cysteines with sites on the target molecule. If this were the case, we might expect this disulfide to be particularly susceptible to reduction. However, the Cys 10–Cys 22 disulfide was the first to be reduced in partial reduction experiments (Fig. 3b). The vicinal disulfide bridge does not, therefore, appear to be unusually reactive. Thus, we speculate that the vicinal disulfide bridge is directly involved in interactions with the target molecule, just as the vicinal disulfide bridge in the AChR appears to be directly involved in acetylcholine binding.
Purification of toxins
Lyophilized crude venom was loaded onto a Vydac C18 analytical rpHPLC column and eluted at flow rate of 1 ml min−1 using a gradient of 5–25% buffer B (0.1% v/v trifluoroacetic acid (TFA) in acetonitrile) over 22 min, followed by a gradient of 25–50% buffer B over 48 min. Buffer A was 0.1% v/v TFA in water. Fractions were collected manually, and individual components were further purified using shallower acetonitrile gradients. Once purified to > 98% homogeneity (as assessed by rpHPLC), peptides were lyophilized and stored at −20 °C until further use. Peptides were pyridethylated as described11 prior to sequencing on an Applied Biosystems 473 protein sequencer.
Peptides were considered insect active if they caused death or prolonged paralysis (>6 h) when injected into mealworms (Tenebrio molitor) and/or house crickets (Acheta domesticus Linnaeus)11. For quantitative analysis of insecticidal activity, the LD50 in A. domesticus was determined as described11.
The effect of J-ACTX-Hv1c on neural transmission was examined using an assay as described17. Briefly, the abdominal and thoracic ganglia were exposed by removal of the dorsal surface and viscera but not the cerci or legs of immobilized P. americana. The toxin was applied directly to the metathoracic ganglion, which was isolated by means of a polyethylene well sealed into place with agar. In this bioassay, stimulation of the insect's posterior sensory organs (cerci) in the absence of toxin elicits reflex movement of the cockroach legs.
Peptide toxins were assayed for their effect in vertebrate smooth (vas deferens) and skeletal (biventer cervicis) nerve muscle tissue as described10. Contractions of each tissue were recorded in the absence of additives or following injection of peptides directly into the bath buffer. δ-ACTX-Hv1 (100 nM), a potent modulator of voltage gated sodium channels38, was used as a positive control.
Vertebrate toxicity assay
Vertebrate toxicity of J-ACTX-Hv1c was determined by subcutaneous injection of toxin in 0.1 ml saline into young BALB/c mice (2.9 ± 0.1 g). Acute toxicity tests were conducted with a minimum number of animals and toxicity was monitored over a period of 48 h.
Bee brain dissociation and electrophysiology.
Neurons were dissociated from the brains of adult European honeybees (Apis mellifera) using a procedure that will be described in detail elsewhere. Briefly, isolated bee brains were incubated for 2 min at room temperature in buffer containing 20 units ml−1 papain. Digestion was terminated with a solution containing 1 mg ml−1 ovomucoid papain/trypsin inhibitor (Type II-O, Sigma) and 1 mg ml−1 bovine serum albumin. Neurons were isolated by gentle trituration though a series of decreasing bore silanized pasteur pipets with fire-polished tips, and then plated on plastic culture dishes coated with poly-d-lysine.
Whole cell voltage clamp recordings were made of bee brain nicotinic AChR and Ca2+, Na+, and K+ channel currents, characterization of which will be reported elsewhere. Neurons were voltage clamped at −90 mV and currents evoked by stepping the membrane potential to −60 mV. Toxin effects on Ca2+ and Na+ currents were tested at the potential with the largest inward current, usually −10 or 0 mV. Toxin effects on K+ channels were determined over a range of membrane potentials (−60 to +60 mV). Nicotinic AChR currents (induced with 30 mM ACh) were obtained at −90 mV. Data were collected and analyzed off line with the PCLAMP and Axotape suite of programs (Axon Instruments).
Folding and purification of synthetic toxins
Synthetic J-ACTX-Hv1c and a Cys 13–Cys 14→Ser-Ser mutant of this toxin were purchased from Auspep. The reduced peptides were purified to homogeneity using rpHPLC, then oxidized/folded in a glutathione redox buffer that promotes disulfide oxidation and shuffling39. At various times after initiation of the folding reaction, aliquots were removed, quenched with HCl, and analyzed using rpHPLC in order to monitor progress of the folding reaction. After 48 h, the reaction mixture was quenched and dialyzed against H2O to remove folding buffer components. The lyophilized dialyzate was then dissolved in H2O and the fully oxidized J-ACTX-Hv1c was purified using rpHPLC.
Chemical confirmation of the vicinal disulfide bond
The disulfide bonding pattern of J-ACTX-Hv1c was determined as described17,18. Briefly, fully oxidized J-ACTX-Hv1c was partially reduced by incubation with a 1:40 molar ratio of Tris-(2-carboxyethyl)-phosphine at pH 3.0 for 15–60 min at either 4 °C, 15 °C, or 37 °C. The reduced cysteines were then carboxamidomethylated by rapid addition of a supersaturated solution of iodoacetamide. The partially reduced/alkylated species were then separated using rpHPLC and their molecular weights determined using mass spectrometry to ascertain the number of cysteines that had been carboxamidomethylated. The peptides were then fully reduced and the unmodified cysteines were pyridethylated. Peptides containing potentially informative combinations of modified cysteine residues were then sequenced on an Applied Biosystems 473 protein sequencer to determine the positions of the carboxamidomethylated and pyridethylated cysteines, from which the positions of disulfide bonds could be inferred.
NMR samples were prepared by dissolving 1.5 mg of J-ACTX-Hv1c or 2.9 mg of the Ser 13-Ser 14 mutant in 250 μl H2O in a susceptibility-matched microcell (Shigemi) and adjusting the pH to 4.95. NMR spectra were recorded at 25 °C and 600 MHz using a 5 mm 1H probe on a Bruker DRX-600 spectrometer. The following homonuclear 2D NMR spectra were recorded for J-ACTX-Hv1c: ECOSY, TOCSY with an MLEV isotropic mixing period of 70 ms, and NOESY with mixing times of 50 and 250 ms. A single NOESY spectrum with a mixing time of 300 ms was recorded for the Ser 13-Ser 14 mutant using a triple resonance inverse probe on a Varian INOVA 600 spectrometer.
Spectra were processed using XWINNMR (Bruker) or Felix97 (Molecular Simulations) and chemical shift assignments were made using XEASY40. Slowly exchanging amides were identified by reconstituting lyophilized J-ACTX-Hv1c in 250 μl 99.96% D2O (Sigma) and immediately recording a time course of 1D spectra for ∼60 min, followed by a time course of eight TOCSY spectra over 16 h.
NOESY crosspeaks were integrated in XEASY and converted to distance restraints (with pseudoatoms where appropriate) using CALIBA41. This yielded 403 nonredundant distance restraints. Twenty dihedral angle restraints27 were derived from 3JHNHα coupling constants measured from either 1D NMR spectra or from inverse Fourier transforms of in-phase NOESY multiplets42. Seven additional φ restraints of −100 ± 80° were applied for residues for which the intraresidue Hα-HN NOE was clearly weaker than that between HN and the Hα of the preceding residue43. The intense intraresidue Hα-HN NOE for Asp 7, combined with a 3JHNHα value of ∼7 Hz, allowed its φ angle to be restrained to 50 ± 40° (refs 31,44).
Stereospecific assignment of β-methylene protons and χ1 restraints were derived for 14 residues using Hα–Hβ coupling constants measured from ECOSY spectra in combination with Hα–Hβ and HN–Hβ NOE intensities45 measured from the 50 ms NOESY experiment. The Hβ protons of Pro 9 and Pro 18 were stereospecifically assigned on the basis of NOE intensities46; this was not possible for Pro 15 and Pro 37 as the β-protons were magnetically equivalent. All four X-Pro peptide bonds were clearly identified as trans on the basis of characteristic NOEs13. Thirteen slowly exchanging amide protons were unambiguously assigned as hydrogen bond acceptors on the basis of preliminary structure calculations. Corresponding hydrogen bond (i-j) restraints of 1.7–2.2 Å and 2.7–3.2 Å were employed for the HNi–Oj and Ni–Oj distances, respectively. The disulfide bonding pattern was determined unequivocally from preliminary structure calculations and distance restraints17 were introduced for the four disulfide bridges in subsequent calculations.
The torsion angle dynamics program DYANA41 was used to calculate 5,000 structures from random starting conformations. The best 100 structures (selected on the basis of final penalty function values) were then refined in X-PLOR47 as described27. The network of experimental restraints was not compatible with either a cis (ω = 0°) or trans (ω = 180°) conformation for the Cys 13–Cys 14 peptide bond. Thus, this ω angle was not restrained in the structure calculations, thereby enabling its value to be determined solely by the network of experimental restraints. The 20 lowest energy X-PLOR structures were used to represent the solution structure of J-ACTX-Hv1c. Molecular graphics analyses were performed using MOLMOL48.
Coordinates and experimental restraints for the ensemble of 20 J-ACTX-Hv1c structures have been deposited in the Protein Data Bank (PDB accession code 1DL0), and 1H chemical shifts have been deposited in BioMagResBank (accession number 4685).
Protein Data Bank
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This work was supported by a grant from the US National Science Foundation to G.F.K. and an Australian Research Council grant to M.J.C. and G.F.K. X.-H.W. and M.C. thank the University of Sydney for award of an Overseas Postgraduate Research Scholarship and Rolf Edgar Lake Postdoctoral Fellowship, respectively. J. Fletcher is gratefully acknowledged for help with structure calculations.
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Wang, Xh., Connor, M., Smith, R. et al. Discovery and characterization of a family of insecticidal neurotoxins with a rare vicinal disulfide bridge. Nat Struct Mol Biol 7, 505–513 (2000). https://doi.org/10.1038/nsb0600_505