A distinct sodium channel voltage-sensor locus determines insect selectivity of the spider toxin Dc1a

Abstract

β-Diguetoxin-Dc1a (Dc1a) is a toxin from the desert bush spider Diguetia canities that incapacitates insects at concentrations that are non-toxic to mammals. Dc1a promotes opening of German cockroach voltage-gated sodium (Na_v) channels (BgNa_v1), whereas human Na_v channels are insensitive. Here, by transplanting commonly targeted S3b-S4 paddle motifs within BgNa_v1 voltage sensors into K_v2.1, we find that Dc1a interacts with the domain II voltage sensor. In contrast, Dc1a has little effect on sodium currents mediated by PaNa_v1 channels from the American cockroach even though their domain II paddle motifs are identical. When exploring regions responsible for PaNa_v1 resistance to Dc1a, we identified two residues within the BgNa_v1 domain II S1–S2 loop that when mutated to their PaNa_v1 counterparts drastically reduce toxin susceptibility. Overall, our results reveal a distinct region within insect Na_v channels that helps determine Dc1a sensitivity, aconcept that will be valuable for the design of insect-selective insecticides.

Introduction

Insect voltage-gated sodium (Na_v) channels share a common architecture with their mammalian orthologues¹. The pore-forming subunit consists of four connected domains (DI-IV), each with six transmembrane segments (S1–S6). These homologous but not identical domains each contain a voltage sensor (S1–S4) and a portion of the pore through which Na⁺ can diffuse (S5–S6)². While they have yet to be identified in insect Na_v channels, each voltage-sensing domain within mammalian and bacterial Na_v channels contains an S3b-S4 helix-turn-helix motif, the voltage-sensor paddle, which drives voltage-sensor activation^{3, 4, 5, 6}. Aside from its vital role in channel gating, the paddle motif is also an important pharmacological target, as peptide toxins interact with this region to modify channel opening^{3, 4, 7}. Insect Na_v channels likely possess similar motifs since residue substitutions in homologous regions can abolish channel susceptibility to toxins found in animal venoms¹. Even though voltage-sensing domains are extensively targeted by naturally occurring peptides, commercially available insecticides such as pyrethroids and oxadiazines typically interact with the channel pore region or intracellular linker between S4 and S5 to disrupt opening or closing (i.e. gating)^{8, 9}. However, insects have responded to this threat by mutating residues at strategic locations within the channel that result in a reduced sensitivity to these compounds⁹. Moreover, the conserved nature of the Na_v channel pore throughout the animal kingdom often leads to undesired biological activity of insecticides in beneficial insect orders or mammals^{8, 9, 10}. Since the amino acid composition of voltage-sensing domains varies considerably between related Na_v-channel isoforms, these regions may replace the pore as a target for designing insecticides with a higher degree of phyletic selectivity, an intriguing notion that has yet to be fully explored.

Na_v channels have been governing electrical excitability in a wide range of organisms for millions of years, even before the development of neurons^{2, 11}. Thus, it is not surprising that ancient organisms such as spiders developed an arsenal of toxins geared towards incapacitating prey by modulating Na_v channel function^{12, 13, 14, 15}. One intriguing specimen is the desert bush spider (Diguetia canities) whose habitat spans the Southwestern deserts of the United States^{16, 17}. The venom of this relatively unexplored primitive weaving spider contains a 57-residue peptide known as DTX9.2 (or β-diguetoxin-Dc1a¹⁸, hereafter Dc1a) (Fig. 1a), one of the most potent insect-selective neurotoxins found in arthropod venoms¹⁹. Although its precise molecular target has yet to be elucidated, neurophysiological studies on housefly larvae revealed excitation of sensory and neuromuscular preparations upon Dc1a application that could be blocked by the Na_v channel-blocking compound tetrodotoxin (TTX)²⁰.

Figure 1. Recombinant production of Dc1a.

a, Primary structure of rDc1a. The non-native N-terminal Ser residue that is a vestige of the TEV protease cleavage site used for recombinant toxin production is highlighted in grey. Disulfide bridge connectivity is shown above the sequence. b, Schematic representation of the pLicC-NSB3 vector used for periplasmic expression of rDc1a. The coding region includes a MalE signal sequence (MalE_SS) for periplasmic export, a His₆ affinity tag, an MBP fusion tag, and a codon-optimized gene encoding rDc1a, with a TEV protease recognition site inserted between the MBP and toxin coding regions. The locations of key elements of the vector are shown, including the ribosome-binding site (RBS). c, SDS-PAGE gels illustrating different steps in the purification of rDc1a. Lanes contain: M, molecular weight markers; lane 1, E. coli cell extract prior to IPTG induction; lane 2, E. coli cell extract after IPTG induction; lane 3, soluble periplasmic extract (the His₆-MBP-rDc1a fusion protein is evident at ~50 kDa); lane 4, Ni-NTA beads after loading the cell lysate; lane 5, eluate-1 from washing Ni-NTA resin with 600 mM imidazole; lane 6, elute-2 from washing Ni-NTA resin with 600 mM imidazole; lane 7, eluate-3 from washing Ni-NTA resin with 600 mM imidazole; lane 8, Ni-NTA beads after elution-3; lane 9, fusion protein sample before TEV protease cleavage; lane 10, fusion protein sample after TEV protease cleavage showing almost complete cleavage of fusion protein to His₆-MBP. d, RP-HPLC chromatogram showing the final step in the purification of rDc1a. The arrow denotes the peak corresponding to correctly folded recombinant rDc1a. Inset is a MALDI-TOF MS spectrum showing the [M+H]⁺ ion for the purified recombinant toxin (obs. = 6484 Da; calc. = 6485.39 Da).

Here, we show that Dc1a potently promotes opening of the German cockroach Na_v channel BgNa_v1²¹ by interacting with the paddle motif in domain II. Surprisingly, Dc1a has little effect on PaNa_v1-expressing neurons isolated from the American cockroach (Periplaneta americana)²², even though the toxin binding site is identical in these two cockroach channels. By exploring regions responsible for the remarkable insect-family specificity of Dc1a, we uncovered an important role for the domain II S1–S2 loop in determining PaNa_v1 resistance to Dc1a. Together with the unique solution structure of Dc1a, our results provide a proof of concept that toxins – and by extension small molecules – can selectively target insect Na_v channels by interacting with their voltage-sensing domains, thus facilitating the development of new insecticides.

Results

Production of recombinant Dc1a

We first produced recombinant Dc1a peptide (rDc1a) using a novel approach that addresses the challenge of correctly folding disulfide-rich spider toxins in E. coli²³. To this end, we created an IPTG-inducible construct (Fig. 1b) that allows for export of a His₆-MBP-Dc1a fusion protein to the E. coli periplasm, where the enzymes involved in disulfide-bond formation are located²³. Upon recovery from the soluble cell fraction, the fusion protein was purified using nickel affinity chromatography (Fig. 1c). Subsequent cleavage and chromatographic purification yielded a single major disulfide-bond isomer with a purity of >98% as assessed by SDS-PAGE and MALDI-TOF mass spectrometry (Fig. 1d). (Note that, with four disulfide bonds, there are 105 possible disulfide-bond isomers of rDc1a.) Overall, the final yield was ~1.1 mg of toxin per litre of culture. Since native Dc1a was unavailable to us, we confirmed the insecticidal activity of rDc1a by injection into the blowfly Lucilia cuprina and the housefly Musca domestica, which yielded LD₅₀ values of 231 ± 32 pmol/g and 493 ± 52 pmol/g, respectively (n = 3) (Supplementary Fig. 1). We therefore conclude that rDc1a has a similar lethality to agricultural pests when compared to native toxin¹⁹.

Solution structure of rDc1a

The development of an efficient bacterial expression system allowed us to produce uniformly ¹³C/¹⁵N-labelled rDc1a protein for structure determination using heteronuclear NMR. ¹H_N, ¹⁵N, ¹³C_α, ¹³C_β, and ¹³C′ resonance assignments for the toxin were obtained from analysis of amide-proton strips in 3D HNCACB, CBCA(CO)NH, and HNCO spectra. Sidechain ¹H and ¹³C chemical shifts were obtained using a 4D HCC(CO)NH-TOCSY experiment, which has the advantage of providing sidechain ¹H-¹³C connectivities²⁴. Complete chemical shift assignments have been deposited in BioMagResBank (Accession Number 19666). CYANA was used for automated NOESY assignment and structure calculation²⁵. The disulfide-bond pattern (1–4, 2–5, 3–8, 6–7) was unambiguously determined from preliminary structures calculated without disulfide-bond restraints²⁶ and is identical to the framework predicted in the UniProtKB entry for Dc1a (P49126). Of the 200 structures that were calculated from random starting conformations, 20 conformers with high stereochemical quality (as judged by MolProbity²⁷) were selected to represent the solution structure of rDc1a (Fig. 2a). Atomic coordinates for the final ensemble of 20 structures are available from the Protein Data Bank (PDB; Accession Number 2MI5).

Figure 2. NMR solution structure of rDc1a.

a, Stereo view of the ensemble of 20 rDc1a structures. The structures are overlaid over the backbone atoms of residues 2–42 and 52–56 in order to highlight the disordered nature of the hairpin loop (residues 43–51, purple) relative to the well-structured ICK region of the toxin. The N- and C-termini are labeled and the β-strands that form the N- and C-terminal β-sheets are colored blue and green, respectively. b, Ribbon representation of rDc1a highlighting the five βstrands (β1–β5) and four disulfide bonds. The three disulfide bonds that form the ICK motif are shown in red while the fourth disulfide that staples the base of the disordered β4–β5 hairpin loop is highlighted in orange. The β-strands and disordered hairpin loop are colored as in panel a. c, Stereo view of the structures of As1a, a typical knottin peptide³⁰, overlaid with rDc1a. The two structures were overlaid for optimal superposition over their core ICK regions (28 residues; region highlighted in cyan) using the CLICK server⁷⁰. Note that the N-terminal β-sheet in rDc1a is quite distinct from the region of structural overlap. d, Electrostatic surface potential of rDc1a with positive and negative charges shown in blue and red, respectively. The molecular orientation is the same as in panels a and c.

Statistics highlighting the high precision and stereochemical quality of the ensemble of rDc1a structures are shown in Supplementary Table 1. The average MolProbity score of 1.24 places the ensemble in the 99th percentile relative to all other structures ranked by MolProbity. The high stereochemical quality of the ensemble stems from a complete absence of bad close contacts and an excellent Ramachandran plot quality (>99% of residues in the most favoured region). The structural ensemble is also highly precise with backbone and heavy-atom RMSD values over all the structural ordered regions (residues 3–42, 52–56) of 0.27 ± 0.07 Å and 0.70 ± 0.11 Å, respectively. The ensemble of rDc1a structures ranks as “very high resolution” based on these measures of precision and stereochemical quality²⁸.

Three of the four disulfide bonds in rDc1a form a classical inhibitor cysteine knot (ICK) motif in which the Cys13–Cys26 and Cys20–Cys40 disulfide bonds and the intervening sections of the polypeptide backbone forming a 23-residue ring that is pierced by the Cys25–Cys54 disulfide bond (Fig. 2b). This ICK motif is commonly found in spider toxins, and this particular scaffold provides these peptides (so-called knottins) with an unusually high degree of chemical, thermal and biological stability¹⁵. However, the structure of rDc1a differs markedly from other ICK toxins, and a DALI search²⁹ of the PDB returned no structural homologs. First, rDc1a contains an additional disulfide bond (Cys42–Cys52) that appears to serve as a molecular staple which limits the flexibility of a disordered serine-rich hairpin loop (residues 43–51) (Fig. 2b). Stapled hairpins of this kind have been observed in only a small number of spider toxins³⁰. Second, the extended N-terminus of rDc1a along with an unusually large loop between Cys26 and Cys40 enables the formation of an N-terminal three-stranded antiparallel β-sheet that is not found in any other knottin (Fig. 2b,c). The uniqueness of the rDc1a structure can readily be seen by comparison with the structure of As1a, a typical spider-venom-derived knottin that modulates the activity of insect Na_V channels³⁰. The two toxins can be aligned over their core ICK regions with an rmsd of 1.9 Å (Fig. 2c) consistent with them both being members of the knottin family. However, when aligned over this ICK region, the N-terminal β-sheet of rDc1a is entirely separate to the region of structural overlap (Fig. 2c). Despite its structural uniqueness, the molecular surface of rDc1a contains a relatively uniform distribution of charged residues (Fig. 2d); moreover, there are no distinct clusters of hydrophobic residues that might mediate an interaction with lipid bilayers^{7, 31} (Supplementary Movie 1).

rDc1a opens the German cockroach Na_v channel BgNa_v1

Diguetia spiders are generalist predators and their diet consists of various insects ranging from small ants to sizeable prey such as grasshoppers and cockroaches^{16, 32}. As such, we decided to test whether rDc1a influences the gating properties of BgNa_v1, a well-studied Na_v channel isoform cloned from the German cockroach^{8, 21}. We functionally expressed BgNa_v1 in Xenopus oocytes and applied various concentrations of rDc1a. At 1 μM, rDc1a produced robust opening of BgNa_v1 at voltages where the channel is normally closed. This was achieved by drastically shifting channel activation to more negative potentials [V_1/2 was shifted from −27.3 ± 0.2 mV (mean ± standard error of the mean – SEM) (slope 4.0 ± 0.2) to −49.4 ± 0.4 mV (slope 3.5 ± 0.3) in the presence of 1 μM rDc1a; Fig. 3a] whereas steady-state inactivation (or channel availability) was only slightly affected (V_1/2 was shifted from −52.2 ± 0.1 mV to −55.1 ± 0.1 mV in the presence of 1 μM rDc1a; Fig. 3a). The rate of recovery from fast inactivation was also not significantly altered (τ is 2.1 ± 0.1 ms vs. 2.2 ± 0.1 ms) (Fig. 3b). Upon addition of 1 μM rDc1a to channels depolarized to −60 mV every 5 s, sodium currents became rapidly visible as a result of the −22 mV shift in voltage-dependent opening of BgNa_v1 (Fig. 3c). Moreover, rDc1a similarly affects the channel when voltage-steps to −60mV were applied every 50 s, suggesting that the toxin can access its binding site when BgNa_v1 is in the resting state¹². Channels completely recovered following washout in toxin-free solution (Fig. 3c). The persistent nature of the emerging current does not stem from the inhibition of fast inactivation by rDc1a since wild-type (WT) BgNa_v1 inherently possesses these characteristics at mildly depolarizing voltages³³. Fitting the Hill equation to the concentration dependence for toxin-induced current potentiation, as determined by shifts in V_1/2, yielded an EC₅₀ of 65 ± 1 nM and a Hill coefficient of 1.19 ± 0.02 (Fig. 3d), suggesting that the toxin may interact with only one binding site.

Figure 3. rDc1a promotes opening of BgNa_v1 channels.

a, Comparison of the gating properties of BgNa_v1 before (black) and after (red) addition of 1 μM rDc1a. Shown are the normalized deduced conductance (G)–voltage (filled circles) (G/G_max) and steady-state inactivation (open circles; I/I_max) relationships. Currents were elicited by 5-mV step depolarizations from a holding voltage of −90 mV or −100 mV, respectively. n = 3–5, and error bars represent SEM. Descriptive values resulting from Boltzmann fits can be found in Results section. b, Recovery from fast inactivation of WT BgNa_v1 before (black) and after (red) addition of 1 μM rDc1a (red) determined by a double-pulse protocol to −30 mV with a varying time between pulses (0–50 ms). Values are reported in the Results section. Data represents n = 4 experiments, and error bars represent the SEM. c, Upon addition of 1 μM rDc1a to channels depolarized to −60 mV every 5 s (red open circles) or 50 s (green open circles) (holding voltage was −100 mV), BgNa_v1-mediated sodium currents become rapidly visible. Channels completely recover after toxin washout. Inset shows current trace before (black) and after (red) addition of 1 μM rDc1a at −60 mV (5 s pulses). Noticeable is the persistent current that appears at this voltage, even in control experiments. Abscissa scale is 10 ms, ordinate scale is 0.3 μA. d, Concentration dependence of rDc1a-induced current potentiation determined from shifts in the midpoint of channel activation (V_1/2). Line represents a fit of the data with the Hill equation resulting in a half maximal concentration (EC₅₀) of 65 ± 1 nM and a slope factor of 1.19 ± 0.02; n = 5 and error bars represent SEM.

Interestingly, 1 μM rDc1a did not affect any of the tested vertebrate Na_v channel isoforms or the hERG channel³⁴, a member of the cardiac voltage-gated potassium (K_v) channel family and an FDA-mandated screening target for potential off-target drug effects (Supplementary Fig. 2a). These results are consistent with the report that Dc1a has no effect in mice when injected intraperitoneally or intracerebroventricularly at doses of 4.1 and 1.0 mg/kg, respectively¹⁹.

rDc1a interacts with the domain II paddle motif in BgNa_v1

Subsequently, we were interested in identifying the receptor sites for rDc1a within insect Na_v channels that result in such a profound shift in the voltage-dependence of activation. Our experiments with BgNa_v1 suggest that rDc1a functions as an excitatory toxin to activate insect Na_v channels at membrane voltages where they are normally closed. Hence, the toxin may interact with S3b-S4 paddle motifs within BgNa_v1 voltage sensors in a manner analogous to previously described β-scorpion toxins^{3, 35, 36}. In order to test this hypothesis, we first needed to establish whether BgNa_v1 contains paddle motifs with similar functions as in K_v channels^{37, 38, 39} and in mammalian Na_v channel isoforms where they have been identified (rNa_v1.2a, rNa_v1.4, rNa_v1.8, rNa_v1.9, and hNa_v1.9)^{3, 4}. To this end, we employed a previously reported approach in which specific S3b-S4 regions from each voltage-sensing domain of a Na_v channel were transplanted into homotetrameric K_v channels (Fig. 4a)^{3, 4, 7}. After several attempts (Fig. 4a), we were able to generate functional chimeras between BgNa_v1 and K_v2.1 using known paddle motif boundaries^{3, 4} (Fig. 4a–b). All of the constructs contain the crucial basic residues that contribute to gating charge movement in K_v channels^{40, 41}, suggesting that each of the four voltage-sensing domains in BgNa_v1 contain paddle motifs that are capable of sensing membrane voltage changes.

Figure 4. Transfer of the voltage sensor paddle motifs from BgNa_v1 to K_v2.1.

a, Sequence alignment of the paddle region of K_v2.1 with the separate S3b–S4 regions of BgNa_v1 arranged per domain. In BgNa_v1, the paddle motifs are not identical and are therefore colored differently: purple, domain I paddle (DI); red, domain II paddle (DII); blue, domain III paddle (DIII); green, domain IV paddle (DIV). In K_v2.1, the paddle motifs are identical and therefore have the same color (black). Functional chimeras are indicated with a green dot in front of the sequence whereas non-functional chimeras are indicated with a red dot. b, and c, Transfer of the BgNa_v1 paddle motifs into K_v2.1. Families of potassium currents (b) and tail current voltage–activation relationships (c) for each chimeric construct (n = 6; error bars represent SEM). Holding voltage was −90 mV for DI, II, and IV, and −120 mV for DIII, and the tail voltage was −60 mV (−100 mV for DIII). Bars in b are 1 μA and 100 ms. d, Kinetics of opening and closing for K_v2.1 channels containing paddle motifs from the four domains of BgNa_v1. Mean time constants (τ) from single-exponential fits to channel activation (filled circles) and deactivation (open circles) are plotted as a function of the voltage at which the current was recorded. n = 6, and error bars represent SEM. e, Potassium currents elicited by depolarizations near the foot (less than 1/3 of maximal activation) of the voltage-activation curve (c) for BgNa_v1/K_v2.1 chimeric constructs. Currents are shown prior to (black) and following (colored) addition of 1 μM rDc1a. rDc1a clearly affects only the DII chimera.

Examination of the conductance-voltage (G-V) relationships for the BgNa_v1/K_v2.1 chimeras reveals that each of the four voltage-sensor paddles has a distinct effect on the gating properties of K_v2.1. This was evidenced by marked differences in the midpoints of activation for the G-V relations for the domain I, II, and IV constructs which are ≥50 mV, 13 ± 2 mV, and −9 ± 2 mV, respectively (Fig. 4c). The G-V relationship for the domain III BgNa_v1/K_v2.1 construct is bimodal, suggesting that the functional coupling between the voltage-sensing domains and the pore has been considerably altered. Although the underlying mechanism is unclear, it is possible that pore opening in this particular chimera may occur when: 1) not all four voltage sensors are fully activated⁴²; or 2) each voltage sensor has transferred only a portion of its charge across the hydrophobic septum^{43, 44, 45}. The latter scenario seems more likely since in the related Shaker K_v channel, all four voltage sensor need to be in the active position before the pore opens^{42, 46}. Moreover, intermediate states have also been observed when measuring gating currents of the Shaker K_v channel in which a mutation immobilized voltage-sensor movement between the resting and activated states, thereby shifting the voltage activation of the ionic currents⁴⁷. Finally, it is worth mentioning that the domain III BgNa_v1/K_v2.1 construct may display multiple open states according to a model that has been described for N-type Ca_v channels⁴⁸.

Remarkably, domain IV paddle motifs from rNa_v1.2a and rNa_v1.4 slow activation kinetics when transplanted into K_v channels. These observations support the notion that the domain IV paddle motif contributes substantially to the overall rate of voltage-sensor activation^{3, 4}. To explore whether the domain IV paddle motif serves a similar role in BgNa_v1, we measured the kinetics of activation and deactivation of the four BgNa_v1 paddle constructs in response to membrane depolarization and repolarization, respectively. Similar to previously studied chimeric channels, the activation and deactivation kinetics observed for the domain IV construct of BgNa_v1/K_v2.1 are slower over a wide voltage range when compared to the other domains (Fig. 4d), suggesting that this particular domain may indeed contribute to fast inactivation in insect Na_v channels^{49, 50, 51}. We obtained additional evidence for the functional role of the domain IV paddle motif in BgNa_v1 gating from experiments with BomIV, a classic insect-selective α-scorpion toxin from the venom of Buthus occitanus mardochei that inhibits fast inactivation in American cockroach neurons⁵². To determine whether BomIV influences BgNa_v1 in a similar fashion, we applied 10 nM to channel-expressing Xenopus oocytes and observed that the toxin indeed slows down fast inactivation (Supplementary Fig. 2b). Subsequent testing on the BgNa_v1/K_v2.1 chimeras revealed that BomIV does not affect WT K_v2.1 or the domain I, II, and III constructs. However, a dramatic toxin-induced inhibition of the domain IV chimera corroborates the importance of this particular voltage-sensing domain in toxin binding and BgNa_v1 fast inactivation (Supplementary Fig. 2b).

We next investigated whether rDc1a modulates the activity of any of the four BgNa_v1/K_v2.1 paddle chimeras and found that 1 μM rDc1a exclusively interacts with the domain II construct whereas domains I, III, IV, and WT K_v2.1 are unaffected (Fig. 4e, Supplementary Fig. 2c). This result suggests that rDc1a specifically targets the domain II voltage sensor within BgNa_v1 to influence channel opening.

rDc1a distinguishes between cockroach Na_v1 channels

Since the domain II paddle motif in BgNa_v1²¹ is identical to that in PaNa_v1²², we expected a similar effect of rDc1a in whole-cell patch-clamp recordings from American cockroach dorsal unpaired median (DUM) neurons. Surprisingly, 1 μM rDc1a had little effect on sodium current amplitude or kinetics in DUM neurons as reflected in the threshold, or V_1/2, of Na_v channel activation or the V_1/2 of steady-state inactivation measurements (Supplementary Fig. 3). Furthermore, there were no significant use-dependent effects on sodium current amplitude or kinetics.

Due to the unexpectedly weak effects of 1 μM rDc1a on voltage-dependent activation of P. americana DUM neurons compared to BgNa_v1 channels from B. germanica, acute toxicity assays were expanded to include both cockroach species (Fig. 5a,b). Remarkably, these bioassays revealed lethal toxicity of rDc1a in B. germanica compared to only mild, reversible effects in P. americana. At doses up to 5 nmol/g, rDc1a generated only minor spastic contractions of the abdomen, some shaking, and evidence of reduced motor activity in P. americana. Even though these effects were completely reversible, a very small percentage of cockroaches did develop paralysis after 24 h (7 ± 7% knockdown, n = 3). One explanation for these observations may be toxin binding to a subset of Na_v channel splice variants⁵³ or the presence of endogenous pharmacologically active auxiliary subunits⁵⁴. Conversely, B. germanica underwent dose-dependent flaccid or contractile paralysis, and lethality was observed after 8 h at all doses tested (Fig. 5a). The KD₅₀ and LD₅₀ values at 24 h post injection were 3.0 ± 1.2 nmol/g and 4.2 ± 3.4 nmol/g (n = 3), respectively. These observations suggest that residues in regions outside of the domain II paddle motif may underlie the remarkably different sensitivity of BgNa_v1 and PaNa_v1 to rDc1a.

Figure 5. Acute toxicity of rDc1a in two cockroach families.

a, Dose–response curve for lethal effects of rDc1a determined 24 h after injection into German cockroaches (B. germanica; family Blattellidae, closed circles). Data were fitted according to Eq. 5 in Materials and Methods and represent mean ± SEM of 3 independent experiments. b, Comparison of the KD₅₀ (open columns) and LD₅₀ (gray columns) doses at 24 h following injection of rDc1a in L. cuprina, M. domestica, and B. germanica. Note: Doses up to 5 nmol/g failed to produce any signs of knockdown or lethality up to 72 h post injection in American cockroaches (Periplaneta americana; family Blattellidae). Data represent mean ± SEM of 3 independent experiments

The S1–S2 loop modulates Na_v channel sensitivity to rDc1a

Sequence alignment of BgNa_v1 with PaNa_v1 reveals a highly conserved S1–S4 voltage sensor in domain II with only three amino acid substitutions (Fig. 6a; Supplementary Fig. 4). Of those, two residues in the S1–S2 loop are potentially accessible for a peptide toxin applied from the extracellular environment (His805 and Asp812 in BgNa_v1 which correspond to a Tyr and Glu in PaNa_v1, respectively). Upon mutating these residues in BgNa_v1 to their PaNa_v1 counterparts, the resulting BgNa_v1^YE construct functionally expressed in Xenopus oocytes with a voltage-dependent activation V_1/2 of −41.5 ± 0.3 mV and a steady-state inactivation V_1/2 of −56.4 ± 0.2 mV (Fig. 6b). Unfortunately, a functional PaNa_v1 clone was unavailable to undertake gain-of-function experiments. Strikingly, 100 nM rDc1a was no longer sufficient to open BgNa_v1^YE whereas this concentration generated large inward sodium currents when applied to WT channels (Fig. 6c, inset). When surveying higher concentrations of rDc1a, it becomes clear that concentrations of more than 1 μM are required to achieve a potentiation with BgNa_v1^YE that is comparable to the WT channel (EC₅₀ ~65 nM) (Fig. 6c). Although we cannot exclude the possibility that rDc1a interacts with other PaNa_v1 domains at such concentrations, administering high doses of the toxin is indeed mildly toxic to P. americana (Fig. 5a,b). Thus, our mutagenesis experiments have uncovered a novel region within insect Na_v channels that helps determine their sensitivity to spider toxins. This locus contributes to the drastically reduced sensitivity of the American cockroach to rDc1a from the venom of the desert bush spider whereas its German counterpart is highly susceptible.

Figure 6. Residues outside of the DII paddle motif contribute to rDc1a binding.

a, Sequence alignment of the DII S1-S2 regions of BgNa_v1 and PaNa_v1 (See Supplementary Fig. 4 for more details). In the extracellular loop connecting S1 to S2, all residues are identical except the H805Y and D812E substitutions (gray background). The T797M substitution within S1 is unlikely to be toxin-accessible. b, Gating properties of the mutant BgNa_v1^YE channel. Shown are the normalized deduced conductance (G)–voltage (filled circles) (G/G_max) and steady-state inactivation (open circles; I/I_max) relationships. Descriptive values can be found in Results section. Currents were elicited by 5 mV step depolarizations from a holding voltage of −90 mV or −100 mV, respectively. n = 3–5, and error bars represent SEM. c, Affinity measure for rDc1a interaction with BgNa_v1^YE (red) compared to WT BgNa_v1 (gray). Concentration dependence for toxin-induced current potentiation (as determined by shifts in V_1/2) is shown. It is clear that alterations in channel opening upon rDc1a exposure are less pronounced with the BgNa_v1^YE mutant when compared to WT BgNa_v1. n = 5 and error bars represent SEM. Inset shows premature channel opening after addition of 100 nM rDc1a to WT BgNa_v1 (grey) whereas BgNa_v1^YE is not yet affected (red). Current traces were evoked by a 50 ms depolarization to −55 mV from a holding potential of −90 mV.

Discussion

The initial goal of this study was to explore the mechanism underlying the efficacy of Dc1a, a potent insecticidal peptide toxin produced by desert bush spiders. To this end, we first produced recombinant rDc1a and determined its solution structure using heteronuclear NMR (Fig. 1 and 2). Next, we tested rDc1a on the heterologously expressed German cockroach channel BgNa_v1 and discovered that the toxin dramatically promotes channel opening (Fig. 3). By taking advantage of the portable nature of S3b–S4 paddle motifs within voltage-sensing domains^{3, 4, 7, 38}, we demonstrated that such motifs also exist in each of the four voltage-sensing domains of BgNa_v1, and that rDc1a selectively interacts with the paddle motif in domain II, a feature that it shares with an extensive list of animal toxins that target this particular paddle motif^{3, 4, 12, 55, 56}, albeit with a high degree of insect selectivity (Fig. 4). Remarkably, our insect assays revealed that German cockroaches are highly susceptible to rDc1a whereas the closely related American cockroach is virtually insensitive despite the fact that the domain II paddle motif in BgNa_v1 is identical to that in PaNa_v1 (Fig. 5 and Supplementary Fig. 4). To elucidate the machinery responsible for this discrepancy, we mutated two residues in the S1–S2 loop that differ between BgNa_v1 and PaNa_v1 (His805 and Asp812) and found that the susceptibility of the resulting BgNa_v1^YE construct to rDc1a is dramatically reduced (Fig. 6). Interestingly, mammalian Na_v channel isoforms possess a Tyr and Ser at the corresponding positions, which presumably contributes to their insensitivity to rDc1a; however, their domain II paddle motif differs as well (Supplementary Fig. 4).

It is interesting to consider potential mechanisms that may underlie the insect-family specificity of rDc1a. For example, the β3–β4 hairpin (Fig. 2b) often houses the pharmacophore in spider ICK toxins that target voltage-gated ion channels⁵⁷, but the unusual architecture of rDc1a (Fig. 2c) and its unique ability to dramatically promote opening of insect Na_v channels (Fig. 3) suggests that it might interact with the voltage sensors of these channels in a unique manner. As such, the domain II S1–S2 loop may play a role in positioning the toxin into a water-filled cavity adjacent to the S3–S4 paddle motif⁵⁸, thereby placing particular toxin residues in strategic positions to stabilize the domain II voltage sensor in an activated position. This hypothesis is strengthened by the observation that the substitution of two residues in the domain II S1–S2 loop within mammalian Na_v1.2 channels decreases sensitivity to the functionally related β-scorpion toxin CssIV by a factor of four⁵⁹. Another possible mechanism is that rDc1a binding to the domain II S1–S2 loop in BgNa_v1 allosterically influences channel opening. Such a hypothesis was postulated when a domain III S1 splice variant of BgNa_v1 revealed an increased susceptibility to the insect-selective β-scorpion toxin Lqh-dprIT₃⁶⁰. However, one caveat is that a depolarizing prepulse is needed to potentiate the maximal effect of CssIV and Lqh-dprIT₃^49,50. Similar to the structurally unrelated spider toxin Magi5 which activates rat Na_v1.2 channels by binding to the domain II voltage sensor⁶¹, rDc1a does not require such a prepulse.

In summary, the current study provides a molecular explanation for the remarkable insect selectivity of Dc1a, the most potent insecticidal toxin identified in the venom of the desert bush spider. Moreover, we have uncovered a novel toxin receptor site within insect Na_v channels that provides a new framework for the design of molecules capable of targeting specific insect families. This knowledge may be used in the future to develop insecticides that target specific insect pests without affecting beneficial insects or endangering human health.

Methods

Chemicals

All chemicals were purchased from Sigma-Aldrich Australia (Castle Hill, NSW, Australia), Sigma-Aldrich USA (St Louis, MO, USA), or Merck Chemicals (Kilsyth, Victoria, Australia) with the exception of isopropyl-β-D-thiogalactopyranoside (IPTG) and streptomycin (Life Technologies, Victoria, Australia), tetrodotoxin (Alomone Labs, Israel), and HPLC-grade acetonitrile (RCI Labscan, Bangkok, Thailand). ¹³C₆-glucose and ¹⁵NH₄Cl were from Sigma-Aldrich Australia. Recombinant His₆-TEV protease (EC 3.4.22.44) was produced in-house using a published protocol⁶².

Production of recombinant rDc1a

A synthetic gene encoding Dc1a, with codons optimised for expression in Escherichia coli, was produced and cloned into a variant of the pLIC-MBP expression vector by GeneArt (Invitrogen, Regensburg, Germany). This vector (pLIC-NSB3) encodes a MalE signal sequence for periplasmic export, a His₆ tag for affinity purification, a maltose binding protein (MBP) fusion tag to aid solubility, and a tobacco etch virus (TEV) protease recognition site directly preceding the codon-optimised Dc1a gene²³. The plasmid encoding Dc1a was transformed into E. coli strain BL21(λDE3) for recombinant toxin production. Protein expression and purification were performed as described previously³⁰ with minor modifications. In summary, cultures were grown in Luria-Bertani broth at 37°C with shaking. Toxin gene expression was induced with 500 μM IPTG at an OD₆₀₀ of 1.0–1.1, then cells were grown at 20 °C for a further 12 h before harvesting by centrifugation for 15 min at 10,500xg For production of uniformly ¹³C/¹⁵N-labelled rDc1a, cultures were grown in minimal medium supplemented with ¹³C₆-glucose and ¹⁵NH₄Cl as the sole carbon and nitrogen sources respectively. The His₆-MBP-toxin fusion protein was extracted from the bacterial periplasm by cell disruption at 27 kPa (TS Series Cell Disrupter, Constant Systems Ltd, Northants, UK), and then captured by passing the extract (buffered in 40 mM Tris, 450 mM NaCl, pH 8.0) over Ni-NTA Superflow resin (Qiagen, Chadstone, Australia). Proteins bound non-specifically were removed by washing with 10 mM imidazole then the fusion protein was eluted with 600 mM imidazole. The eluted fusion protein was concentrated to 10 ml and the buffer was exchanged to remove imidazole. Reduced and oxidised glutathione were then added to a final concentration of 0.6 mM and 0.4 mM, respectively, to maintain TEV protease activity and promote folding of the protein. Approximately 100 μg of His₆-tagged TEV protease was added per mg of rDc1a, then the cleavage reaction was allowed to proceed at room temperature for 12 h. The cleaved His₆-MBP and His₆-TEV were precipitated by addition of 1% trifluoroacetic acid (TFA), then the sample was centrifuged at 16,000xg. The supernatant was filtered using a 0.45 μm syringe filter (Millipore, MA, USA) and subjected to further purification using RP-HPLC. RP-HPLC was performed on a Vydac C18 column (250 × 4.6 mm, particle size 5 μm) using a flow rate of 1 ml/min and a gradient of 20–45% Solvent B (0.043% trifluoroacetic acid (TFA) in 90% acetonitrile) in Solvent A (0.05% TFA in water) over 30 min. rDc1a contains a non-native N-terminal serine residue (a vestige of the TEV protease cleavage site), making it one-residue longer than native Dc1a.

Mass spectrometry

Toxin masses were confirmed by matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF MS) using a Model 4700 Proteomics Bioanalyser (Applied Biosystems, CA, USA). RP-HPLC fractions were mixed (1:1 v:v) with α-cyano-4 hydroxycinnamic acid matrix (5 mg/ml in 50/50 acetonitrile/H₂O) and MALDI-TOF spectra were acquired in positive reflector mode. All reported masses are for monoisotopic [M + H]⁺ ions.

Structure determination

Several buffer conditions were screened to optimize quality of NMR data recorded for rDc1a, including 20 mM phosphate buffer, pH 6.0; 20 mM MES buffer, pH 6.0 and 20 mM acetate buffer pH 5.0. Initial buffer screening revealed the acetate buffer, pH 5.0 was optimal for acquisition of NMR data. Recombinant ¹⁵N/¹³C-labelled rDc1a was dissolved in 20 mM acetate buffer, pH 5.0 to a final concentration of 350 μM. 5% ²H₂O was added, then the sample was filtered using a low-protein-binding Ultrafree-MC centrifugal filter (0.22 μm pore size; Millipore, MA, USA) and 300 μL was added to a susceptibility matched 5 mm outer-diameter microtube (Shigemi Inc., Japan). NMR data were acquired at 25°C using a 900 MHz NMR spectrometer (Bruker BioSpin, Germany) equipped with a cryogenically cooled probe. 3D and 4D data used for resonance assignments were acquired using non-uniform sampling (NUS). Sampling schedules that approximated the signal decay in each indirect dimension were generated using sched3D²⁶. NUS data were processed using the Rowland NMR toolkit (www.rowland.org/rnmrtk/toolkit.html) and maximum entropy parameters were automatically selected as previously described^{63. 13}C- and ¹⁵N-edited HSQC-NOESY (mixing time of 200 ms) experiments were acquired using uniform sampling. All experiments were acquired in H₂O containing 5% ²H₂O. Dihedral angles (φ, ψ) were derived from TALOS+ chemical shift analysis⁶⁴ and the restraint range for structure calculations was set to twice the estimated standard deviation. The Gly9–Pro10 and Arg36–Pro37 peptide bond was determined to be in the trans conformation on the basis of characteristic NOEs and the C_α and C_β chemical shifts of Pro residues. NOESY spectra were manually peak picked and integrated, then peak lists were automatically assigned, distance restraints extracted, and an ensemble of structures calculated using the torsion angle dynamics package CYANA 3.0²⁵. The tolerances used for CYANA 3.0 were 0.025 ppm in the direct ¹H dimension, 0.03 ppm in the indirect ¹H dimension, and 0.3 ppm for the heteronucleus (¹³C/¹⁵N). During the automated NOESY assignment/structure calculation process, CYANA assigned 92% of all NOESY crosspeaks (2271 out of 2469).

Insecticidal assays

rDc1a was dissolved in insect saline and injected into the ventro-lateral thoracic region of adult sheep blowflies (Lucilia cuprina;), adult houseflies (M. domestica), adult American cockroaches (Periplaneta americana) and adult German cockroaches (Blatella germanica)⁶⁵. Injections were made using a 1.0 ml Terumo Insulin syringe (B-D Ultra-Fine, Terumo Medical Corporation, Maryland, USA) with a fixed 29 G needle fitted to an Arnold hand micro-applicator (Burkard Manufacturing Co. Ltd., England). A maximum volume of 2 μl was injected per L. cuprina, M. domestica and B. germanica and 5 μl for the larger P. americana. Thereafter, flies were individually housed in 2-ml tubes and provided with 10% sucrose while cockroaches were housed in closed Petri dishes and provided with dry food and water. The paralytic activity was then determined over a 24 h period. For each acute toxicity assay up to fourteen doses of rDc1a (n = 10 insects per dose) and the appropriate control (insect saline; n = 10 insects) were used. The assay was repeated three times. Median knockdown (KD₅₀) and median lethal (LD₅₀) doses were calculated as described previously⁶⁶ and averaged to produce KD₅₀ and LD₅₀ values.

Electrophysiological measurements on DUM neurons

DUM neurons were isolated from unsexed adult American cockroaches (P. americana) as described previously³⁰. Briefly, terminal abdominal ganglia were removed and placed in normal insect saline (NIS) containing (in mM): NaCl 180, KCl 3.1, N-hydroxyethylpiperazine-N-ethanesulfonic acid (HEPES) 10 and D-glucose 20. Ganglia were then incubated in 1 mg/ml collagenase (type IA) for 40 min at 29°C. Following enzymatic treatment, ganglia were washed three times in NIS and triturated through a fire-polished Pasteur pipette. The resultant cell suspension was then distributed onto 12-mm diameter glass coverslips pre-coated with 2 mg/ml concanavalin A (type IV). DUM neurons were maintained in NIS supplemented with 5 mM CaCl₂, 4 mM MgCl₂, 5% foetal bovine serum and 1% penicillin and streptomycin (Life Technologies, Victoria, Australia), and maintained at 29 °C, 100 % humidity. Ionic currents were recorded in voltage-clamp mode using the whole-cell patch-clamp technique employing version 10.2 of the pCLAMP data acquisition system (Molecular Devices, Sunnyvale, CA). Data were filtered at 5–10 kHz with a low-pass Bessel filter with leakage and capacitive currents subtracted using −P/4 procedures. Digital sampling rates were set between 15 and 25 kHz depending on the length of the protocol. Single-use 0.8–2.5 MΩ electrodes were pulled from borosilicate glass and fire-polished prior to current recordings. Liquid junction potentials were calculated using JPCALC, and all data were compensated for these values. Cells were bathed in external solution through a continuous pressurized perfusion system at 1 ml/min, while toxin solutions were introduced via direct pressurized application via a perfusion needle at ~50 μl/min (Automate Scientific, San Francisco, CA). Control data were not acquired until at least 20 min after whole-cell configuration was achieved to eliminate the influence of fast time-dependent shifts in steady-state inactivation resulting in Na_V channel current (I_Na) rundown. All experiments were performed at ambient room temperature (20–23°C). To record I_Na, the external bath solution contained (in mM): NaCl 80, CsCl 5, CaCl₂ 1.8, tetraethylammonium chloride 50, 4-aminopyridine 5, HEPES 10, NiCl₂ 0.1, and CdCl₂ 1, adjusted to pH 7.4 with 1 M NaOH. The pipette solution contained (in mM): NaCl 34, CsF 135, MgCl₂ 1, HEPES 10, ethylene glycol-bis(2-aminoethylether)-N,N,N′,N′-tetraacetic acid 5, and ATP-Na₂ 3, adjusted to pH 7.4 with 1 M CsOH. To eliminate any influence of differences in osmotic pressure, all internal and external solutions were adjusted with sucrose to 400 ± 5 mOsmol/l. Experiments were rejected if there were large leak currents or currents showed signs of poor space clamping.

Off-line data analysis was performed using Axograph X version 1.3 or Clampfit 10 (Molecular Devices, USA). Voltage-activation relationships were obtained by measuring steady-state currents elicited by stepwise depolarizations of 5–10 mV from a holding potential of −90 mV and calculating peak conductance (G_Na) using the following equation: $G=I_{\text{Na}}/(V_{\mathrm{m}}-E_{\text{rev}})$ where G is peak conductance, I_Na is peak inward sodium current, V_m is the test potential, and E_rev is the reversal potential. Reversal potentials were individually estimated for each data set⁶⁷ by fitting the I_Na-V data with the following equation: $I_{\text{Na}}={[1+exp(-0.03937\times \mathrm{z}\times (V_{\mathrm{m}}-{\mathrm{V}}_{1/2}))]}^{-1}\times g\times (V_{\mathrm{m}}-V_{\text{rev}})$ where z is the apparent gating charge, g is a factor related to the number of channels contributing to the macroscopic whole-cell I_Na, V_m is equal to the voltage potential of the pulse, and V_1/2 is the voltage at half-maximal activation. The normalized conductance was fitted to a two-state Boltzmann function of the form: G/G_max = 1/(1+exp((−0.3937 ×z ×(V_m–V_1/2))) where G, z, V_m and V_1/2 have the same meaning. The voltage dependence of steady-state Na_v channel inactivation (h_∞/V) data were normalized to the maximum peak current in the control or maximum peak current and fitted using the following Boltzmann equation: ${\mathrm{h}}_{\infty }=\mathrm{A}/(1+exp((V_{\mathrm{m}}-V_{12})/k))$ where A is the fraction of control maximal peak I_Na (value of 1.0 under control conditions), V_1/2 is the midpoint of steady-state inactivation, k is the slope factor, and V_m is the prepulse voltage. The rate of recovery from fast inactivation data were fitted to the following single exponential function: I/I_max = 1-exp(−τ/t) where τ is the time constant of current recovery. Dose–response curves to determine LD₅₀ and KD₅₀ values were fitted using the following form of the logistic equation: $y=1/{(1+[x]/{\text{Dose}}_{50})}^{n\mathrm{H}}$ where x is the toxin dose, n_H the Hill coefficient (slope parameter), and Dose₅₀ is the median inhibitory dose causing lethality (LD₅₀) or knockdown (KD₅₀). Non-linear curve-fitting of data were performed using GraphPad Prism version 6.00c for Macintosh (GraphPad Software, San Diego) or Origin 8 (OriginLab, MA, USA). Comparisons of two sample means were made using a paired Student’s t-test and differences were considered to be significant if p < 0.05. All data are presented as mean ± standard error of the mean (SEM) of n independent experiments.

Electrophysiological measurements on oocytes

Chimeras and channel mutants were generated using sequential polymerase chain reactions (PCRs) with K_v2.1Δ7⁶⁸ and BgNa_v1²¹ as templates. The Kv2.1Δ7 construct contains seven point mutations in the outer vestibule, rendering the channel sensitive to agitoxin-2. The DNA sequence of all constructs and mutants was confirmed by automated DNA sequencing and cRNA was synthesized using T7 polymerase (mMessage mMachine kit, Life Technologies, USA) after linearizing the DNA with appropriate restriction enzymes. Channel constructs including hNa_v1.1–hNa_v1.7 and hERG were expressed in Xenopus oocytes by themselves or, in the case of BgNa_v1, together with the TipE subunit⁶⁹ (1:5 molar ratio), and studied following 1–2 days incubation after cRNA injection (incubated at 17°C in 96mM NaCl, 2mM KCl, 5mM HEPES, 1mM MgCl₂ and 1.8mM CaCl₂, 50μg/ml gentamycin, pH 7.6 with NaOH) using two-electrode voltage-clamp recording techniques (OC-725C, Warner Instruments, USA) with a 150-μl recording chamber. Data were filtered at 4 kHz and digitized at 20 kHz using pClamp software (Molecular Devices, USA). Microelectrode resistances were 0.5–1 MΩ when filled with 3M KCl. For most K_v channel experiments, the external recording solution contained (in mM): 50 KCl, 50 NaCl, 5 HEPES, 1 MgCl₂, 0.3 CaCl₂, pH 7.6 with NaOH. For Na_v channel experiments, the external recording solution contained (in mM): 96 NaCl, 2 KCl, 5 HEPES, 1 MgCl₂ and 1.8 CaCl₂, pH 7.6 with NaOH. All experiments were performed at room temperature (~22 °C). Leak and background conductances, identified by blocking the channel with agitoxin-2 (Alomone labs) or tetrodotoxin (TTX), were subtracted for most of the K_v or BgNa_v1 currents shown.

Voltage–activation relationships were obtained by measuring tail currents for K_v channels or steady-state currents and calculating conductance for Na_v channels. Protocols for other measurements are described in the figure legends. After addition of the toxin to the recording chamber, the equilibration between the toxin and the channel was monitored using weak depolarizations elicited at 5-s intervals. For all channels, we recorded voltage–activation relationships in the absence and presence of toxin. Off-line data analysis was performed using Clampfit 10 (Molecular Devices, USA), Origin 8.0 (Originlab) and Microsoft Solver (Microsoft Excel).