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Research ArticleArticle

Identification and Characterization of ProTx-III [μ-TRTX-Tp1a], a New Voltage-Gated Sodium Channel Inhibitor from Venom of the Tarantula Thrixopelma pruriens

Fernanda C. Cardoso, Zoltan Dekan, K. Johan Rosengren, Andelain Erickson, Irina Vetter, Jennifer R. Deuis, Volker Herzig, Paul F. Alewood, Glenn F. King and Richard J. Lewis
Molecular Pharmacology August 2015, 88 (2) 291-303; DOI: https://doi.org/10.1124/mol.115.098178

Abstract

Spider venoms are a rich source of ion channel modulators with therapeutic potential. Given the analgesic potential of subtype-selective inhibitors of voltage-gated sodium (NaV) channels, we screened spider venoms for inhibitors of human NaV1.7 (hNaV1.7) using a high-throughput fluorescent assay. Here, we describe the discovery of a novel NaV1.7 inhibitor, μ-TRTX-Tp1a (Tp1a), isolated from the venom of the Peruvian green-velvet tarantula Thrixopelma pruriens. Recombinant and synthetic forms of this 33-residue peptide preferentially inhibited hNaV1.7 > hNaV1.6 > hNaV1.2 > hNaV1.1 > hNaV1.3 channels in fluorescent assays. NaV1.7 inhibition was diminished (IC50 11.5 nM) and the association rate decreased for the C-terminal acid form of Tp1a compared with the native amidated form (IC50 2.1 nM), suggesting that the peptide C terminus contributes to its interaction with hNaV1.7. Tp1a had no effect on human voltage-gated calcium channels or nicotinic acetylcholine receptors at 5 μM. Unlike most spider toxins that modulate NaV channels, Tp1a inhibited hNaV1.7 without significantly altering the voltage dependence of activation or inactivation. Tp1a proved to be analgesic by reversing spontaneous pain induced in mice by intraplantar injection in OD1, a scorpion toxin that potentiates hNaV1.7. The structure of Tp1a as determined using NMR spectroscopy revealed a classic inhibitor cystine knot (ICK) motif. The molecular surface of Tp1a presents a hydrophobic patch surrounded by positively charged residues, with subtle differences from other ICK spider toxins that might contribute to its different pharmacological profile. Tp1a may help guide the development of more selective and potent hNaV1.7 inhibitors for treatment of chronic pain.

Introduction

Voltage-gated sodium (NaV) channels are transmembrane proteins that underlie action potentials in excitable cells where they contribute to a broad range of vital biologic processes (Diss et al., 2004). The NaV channel superfamily comprises nine subtypes (NaV1.1–NaV1.9) that differ in their primary structure as well as in pharmacological and functional properties. Different subtypes are involved in specialized functions depending on their distribution, biophysical properties, and density (Catterall, 2012). These differences provide an opportunity for tissue-specific inhibition of one subtype without affecting the function of others.

Aberrant function and expression of NaV channels are associated with a range of complex pathologic conditions, such as chronic pain, epilepsy, and cardiac arrhythmias (Rogers et al., 2006; Catterall et al., 2010; Remme and Bezzina, 2010). Remarkably, individuals with loss-of-function mutations in the gene encoding the pore-forming α subunit of human NaV1.7 (hNaV1.7) have a complete insensitivity to pain (Cox et al., 2006), with no other sensory deficits except a lack of smell (anosmia) due to the role of hNaV1.7 in olfaction (Weiss et al., 2011; Rupasinghe et al., 2012). Thus, hNaV1.7 has become an exciting, genetically validated target for treating pain disorders (Liu and Wood, 2011; King and Vetter, 2014; Minett et al., 2014).

Spider venoms are a rich source of disulfide-rich peptides with therapeutic potential (Saez et al., 2010; King, 2011). Most of these peptides are ion channel modulators that target a wide range of prey species from invertebrates to mammals (King and Hardy, 2013; Smith et al., 2013). It was recently shown that spider venoms contain at least 12 discrete classes of NaV channel spider toxins (NaSpTxs) (Klint et al., 2012), and hence, we decided to screen these venoms for specific inhibitors of hNaV1.7 by optimizing a high-throughput fluorescent imaging plate reader (FLIPR)–based screen we recently described for discovering modulators of endogenously expressed hNaV1.7 (Vetter et al., 2012; Klint et al., 2015). By selectively activating hNaV1.7 using the small-molecule agonist veratridine in combination with the NaV1.6/NaV1.7-selective potentiator OD1, a scorpion-venom peptide (Jalali et al., 2005; Maertens et al., 2006; Vetter et al., 2012; Durek et al., 2013), we were able to directly screen for hNaV1.7 inhibitors. This assay is coupled to calcium influx, allowing high-throughput screens for molecules that inhibit hNaV1.7 using calcium-sensitive fluorescent dyes.

Here, we describe the identification of a potent hNaV1.7 inhibitor from the venom of the tarantula Thrixopelma pruriens. This 33-residue peptide, named μ-TRTX-Tp1a (hereafter, Tp1a) on the basis of the rational nomenclature for spider toxins (King et al., 2008), was produced by bacterial expression and chemical synthesis, and its NaV subtype selectivity was determined using fluorescent imaging and electrophysiology. We show that Tp1a is analgesic in vivo and reveal its three-dimensional (3D) structure as determined using NMR.

Materials and Methods

Animals.

For behavioral assessment, adult male C57BL/6J mice aged 6–8 weeks and weighing 20–25 g were used. Mice were housed in groups of three or four per cage, under a 12-hour light-dark cycle, with standard rodent chow and water provided ad libitum.

Ethics Statement.

Ethical approval for in vivo experiments was obtained from The University of Queensland animal ethics committee. Experiments involving animals were conducted in accordance with the Animal Care and Protection Regulation Qld (2012), the Australian Code of Practice for the Care and Use of Animals for Scientific Purposes, 8th edition (2013), and the International Association for the Study of Pain Guidelines for the Use of Animals in Research.

Spider Venoms.

Venoms obtained from 40 species of theraphosid spiders by cheliceral electrostimulation were kept frozen at –20°C before being lyophilized (Herzig and Hodgson, 2009).

Cell Culture.

Cell culture reagents were from Gibco (Life Technologies Corporation, Carlsbad, CA) unless otherwise stated. The human neuroblastoma cell line SH-SY5Y was maintained at 37°C in a humidified 5% CO2 incubator in RPMI medium supplemented with 15% fetal bovine serum (FBS) and 2 mM l-glutamine. Chinese hamster ovary (CHO) cells expressing recombinant hNaV channels (EZ cells; ChanTest Corp., Cleveland, OH) were maintained at 37°C in a humidified 5% CO2 incubator in F-12 medium supplemented with 10% FBS, 100 U/ml penicillin, and 100 μg/ml streptomycin. Human embryonic kidney 293 (HEK 293) cells expressing recombinant hNaV subtypes coexpressed with the β1 subunit (SB Drug Discovery, Glasgow, UK) were maintained at 37°C in a humidified 5% CO2 incubator in minimal essential medium (Sigma-Aldrich, St. Louis, MO) supplemented with 10% FBS, 100 U/ml penicillin and 100 μg/ml streptomycin, 2 mM l-glutamine, and variable concentrations of blasticidin, geneticin, and zeocin, according to manufacturer’s protocol. A tetrodotoxin-resistant rat NaV1.6 cell line was provided by Steven Waxman (Yale School of Medicine, New Haven, CT) and cultured in Dulbecco’s modified Eagle’s medium supplemented with 10% FBS and 500 μg/ml geneticin. Replicating cells were subcultured every 3–4 days in a 1:5 ratio using 0.25% trypsin/EDTA.

Venom Screens against hNaV1.7.

Venoms previously identified to inhibit SH-SY5Y cell responses stimulated nonspecifically with veratridine (Klint et al., 2015) were rescreened for hNaV1.7 inhibition as previously described (Vetter et al., 2012). In brief, SH-SY5Y cells were plated at 40,000 cells per well in 384-well flat clear-bottom black plates (Corning Inc., Corning, NY) and cultured at 37°C in a humidified 5% CO2 incubator for 48 hours. Cells were loaded with 20 μl/well Calcium 4 dye (Molecular Devices, Sunnyvale, CA) reconstituted in assay buffer containing (in millimolar) 140 NaCl, 11.5 glucose, 5.9 KCl, 1.4 MgCl2, 1.2 NaH2PO4, 5 NaHCO3, 1.8 CaCl2, and 10 HEPES (pH 7.4) and incubated for 30 minutes at 37°C in a humidified 5% CO2 incubator. Fluorescence responses were recorded using excitation of 470–495 nm and emission of 515–575 nm for 10 seconds to set the baseline, then again 600 seconds after addition of 10, 1, or 0.1 μg of venom/well and for a further 300 seconds after coaddition of 3 μM veratridine and 30 nM OD1.

Venom Peptide Purification.

Venom from T. pruriens (1 mg) was dissolved in 100 μl of Milli-Q water containing 0.05% trifluoroacetic acid (TFA) (Auspep, Tullamarine, VIC, Australia) and 5% acetonitrile (ACN) (Sigma-Aldrich) and centrifuged at 14,000 rpm for 10 minutes to remove particulates. Venom was fractionated on a C18 reversed-phase (RP) high-performance liquid chromatography (HPLC) column (Jupiter 250 × 4.6 mm, 5 μm; Phenomenex, Torrance, CA) using a gradient established in Milli-Q water containing 0.05% TFA (solvent A) and 90% ACN in 0.043% TFA (solvent B). Fractionation was achieved using the following gradient: 5% B for 5 minutes, followed by 5–20% B over 5 minutes, then 20–50% B over 70 minutes at a flow rate of 1 ml/min. Peaks were collected by monitoring the absorbance at 214 nm and fractions lyophilized before storage at –20°C.

Mass Spectrometry and Sequencing.

Peptide masses were determined by matrix-assisted laser desorption/ionization time of flight (TOF) mass spectrometry (MS) using a 4700 Proteomics Bioanalyzer model (Applied Biosystems, Carlsbad, CA). Peptides dissolved in water were mixed 1:1 (v/v) with α-cyano-4-hydroxy-cinnamic acid matrix (7 mg/ml in 50% acetonitrile) and mass spectra acquired in positive reflector mode. Reported masses are for the monoisotopic M+H+ ions. C-terminal amidation was determined by electrospray mass spectrometry performed using a Triple TOF 5600 system (AB Sciex, Framingham, MA) coupled to a DGU-20AD Nano HPLC (Shimadzu, Kyoto, Japan). For analysis, native Tp1a was reconstituted in 0.1% formic acid and loaded into the HPLC, and data were analyzed using Analyst TF 1.6 software (AB Sciex). N-terminal sequencing was performed by the Australian Proteome Analysis Facility (Sydney, NSW, Australia). In brief, for N-terminal sequencing, the peptide was dissolved in urea (4 M) in ammonium bicarbonate (50 mM) and reduced with dithiothreitol (100 mM) at 56°C for 1 hour under argon. The sample was then alkylated using acrylamide (220 mM) for 0.5 hour in the dark. The reaction was quenched by the addition of excess dithiothreitol. After desalting by RP-HPLC, the collected fraction was loaded onto precycled bioprene discs and subjected to 31 cycles of Edman N-terminal sequencing using a 494 Procise Protein Sequencing System (Applied Biosystems). For amino acid analysis, samples were reconstituted in 150 μl of 20% acetonitrile/0.1% TFA and dried for hydrolysis. For high-sensitivity amino acid analysis, samples underwent 24-hour gas phase hydrolysis in 6 M HCl at 110° C. The amount of Asn/Asp and Gln/Glu is reported as the sum of the related amino acids. Cysteine and tryptophan are not analyzed using this method. After hydrolysis, amino acids were analyzed using the Water AccQTag Ultra chemistry on a Waters Acquity UPLC (Waters Corporation, Milford, MA).

Recombinant Production.

A synthetic gene encoding Tp1a, with codons optimized for expression in Escherichia coli, was produced and cloned into the pLicC vector by GeneArt (Life Technologies). This vector contains a MalE signal sequence for periplasmic expression, a His6 tag for nickel affinity purification, a maltose binding protein (MBP) tag to enhance solubility, and a tobacco etch virus (TEV) protease recognition site preceding the Tp1a gene (Klint et al., 2013). The expression plasmid was transformed in E. coli strain BL21(DE3) and cells cultured in Luria-Bertani medium at 37°C, 180 rpm. Expression of Tp1a was induced with 0.5 mM isopropyl β-d-1-thiogalactopyranoside at an Optical Density600 of 1.2, and cells were cultured for further 16 hours at 16°C. Cells were harvested by centrifugation for 20 minutes at 10,000 rpm, then the pellet was resuspended in FastBreak lysis buffer (Promega, Madison, WI) containing 0.2 mg/ml lysozyme and 10 U/ml DNase. After 30-minute incubation at room temperature, cells were centrifuged for 30 minutes at 12,000 rpm and the supernatant collected. The fusion protein was captured by passing the lysate through a Ni–nitrilotriacetic acid resin (Sigma-Aldrich) followed by washing with 15 mM imidazole in tris NaCl (TN) buffer (25 mM Tris, 150 mM NaCl, pH 8). The fusion protein was eluted with 500 mM imidazole in TN buffer, then the eluate was concentrated and desalted against TN buffer using a 30-kDa cut-off Amicon Ultra-15 centrifugal filter (Millipore, Billerica, MA). Reduced (0.6 mM) and oxidized (0.4 mM) glutathione were added to the sample to facilitate TEV protease cleavage performed with 0.02 mg/ml TEV protease at 30°C for 16 hours. The sample was then filtered using a 30-kDa cut-off Amicon Ultra-15 centrifugal filter to remove MBP and uncleaved fusion protein, and then the eluate was loaded onto a C18 column (Vydac 4.6 × 250 mm, 5 μm; Grace Discovery Sciences, Columbia, MD). RP-HPLC was performed with an Ultimate 3000 LC system (Dionex, Sunnyvale, CA) using a gradient in Milli-Q water/0.05% TFA (solvent A) and 90% ACN/0.045% TFA (solvent B). Peaks were collected, lyophilized, and stored at –20°C.

Peptide Synthesis.

Both the C-terminal amide (Tp1a-NH2) and acid (Tp1a-OH) forms of Tp1a were assembled on a Symphony automated peptide synthesizer (Protein Technologies Inc., Tucson, AZ) using Rink-amide (loading 0.64 mmol/g) or Fmoc-Leu-Wang (loading 0.56 mmol/g) polystyrene resins, respectively, on a 0.1 mmol scale. Fmoc deprotection was achieved using 30% piperidine/dimethylformamide. Couplings were performed in dimethylformamide using five equivalents of Fmoc–amino acid/2-(1H-benzotriazol-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate/N,N,-diisopropylethylamine (1:1:1) relative to resin loading for 2 × 20 minutes. Amino acid side chains were protected as follows: Arg(2,2,4,6,7-pentamethyldihydrobenzofuran-5-sulfonyl) Asn(triphenylmethyl [Trt]), Asp(tert-butyl ester), Cys(Trt), Glu(tert-butyl ester), His(Trt), Lys(tert-butoxycarbonyl), Ser(tert-butyl), Trp(tert-butoxycarbonyl). Cleavage from resin and removal of side chain protecting groups was achieved by treatment with 95% TFA/2.5% triisopropylsilane/2.5% H2O at room temperature for 2 hours. After most of the cleavage solution was evaporated under a stream of N2, the products were precipitated and washed with cold Et2O and lyophilized from 50% MeCN/0.1% TFA/H2O [Tp1a-NH2: 230 mg; electrospray ionization (ESI)–MS (m/z): calculated (average) [calc. (avg.)] 762.7 [M+5H]5+, found 762.6; Tp1a-OH: 289 mg; ESI-MS (m/z): calc. (avg) 762.9 [M+5H]5+, found 762.8]. The crude products were purified by preparative HPLC to give 108 and 90 mg of reduced Tp1a-NH2 and Tp1a-OH, respectively.

Oxidative Folding.

Purified reduced peptide (50 mg of Tp1a-NH2 or 45 mg of Tp1a-OH), reduced glutathione (100 equivalent), and oxidized glutathione (10 equivalent) were dissolved in 6 M GnHCl (21 ml) then added to a solution of 0.36 M NH4OAc (pH 8.0, 230 ml) and stirred at room temperature with exposure to air for 48 hours. The single major products were isolated by preparative HPLC [Tp1a-NH2: 30 mg; ESI-MS (m/z): calc. (avg) 761.5 [M+5H]5+, found 761.5; Tp1a-OH: 24.5 mg; ESI-MS (m/z): calc. (avg) 761.7 [M+5H]5+, found 761.5].

Membrane Potential Assay.

Changes in membrane potential were measured in cells expressing human NaV subtypes on a FLIPRTETRA using membrane potential dye red (Molecular Devices). The subtypes hNaV1.1/β1, hNaV1.2/β1, hNaV1.3/β1, hNaV1.4/β1, hNaV1.5/β1, rNaV1.6, hNaV1.7/β1, and hNaV1.8/β1 stably expressed in HEK cells (SB Drug Discovery) were plated at 10,000 cells per well in 384-well flat clear-bottom black plates (Corning) and cultured in complete media at 37°C in a humidified 5% CO2 incubator for 24 hours. For assays, the medium was removed and cells were loaded with 20 μl/well of membrane potential dye red reconstituted in assay buffer containing (in millimolar) 140 NaCl, 11.5 glucose, 5.9 KCl, 1.4 MgCl2, 1.2 NaH2PO4, 5 NaHCO3, 1.8 CaCl2, and 10 HEPES (pH 7.4). Tp1a was added to cells and incubated for 30 minutes at 37°C in a humidified 5% CO2 incubator. Changes in membrane potential were recorded using excitation (510–545 nm) and emission (565–625 nm) for 10 seconds to set the baseline, then for a further 300 seconds after addition of 50–70 μM veratridine.

Patch-Clamp Electrophysiology.

Experiments were recorded in recombinant CHO (ChanTest) or HEK 293 (SB Drug Discovery) cells expressing specific NaV subtypes, and Na+ currents were measured using an automated whole-cell patch-clamp electrophysiology system (QPatch 16X; Sophion, Ballerup, Denmark). The extracellular solution comprised (in millimolar) 1 CaCl2, 1 MgCl2, 5 HEPES, 3 KCl, 140 NaCl, 0.1 CdCl2, 20 TEA-Cl at pH 7.3, and 320 mOsm, and the intracellular solution comprised (in millimolar) 140 CsF, 1/5 EGTA/CsOH, 10 HEPES, 10 NaCl at pH 7.3, and 320 mOsm. The elicited currents were sampled at 25 kHz and filtered at 4 kHz. The average seal, whole-cell, and chip resistance values were (in megaohms) 1700, 1672, and 2.1 for CHO cells and 3690, 997, and 2.08 for HEK cells, respectively. Cells were maintained at a holding potential of −80 mV and Na+ currents elicited by 20-millisecond voltage steps to 0 mV from a −120-mV conditioning pulse applied for 200 milliseconds. For dose-response experiments, cells were incubated for 5 minutes with increasing concentrations of Tp1a. For on-rate measurements, Na+ currents were assessed at 15-second intervals immediately after addition of Tp1a. For off-rate measurements, cells were incubated for 10 minutes with Tp1a and Na+ currents assessed at 5-minute intervals during saline washes. The Kon, Koff, and Kd were calculated using Kd = Koff/Kon (nanomolar), where Koff = 1/τoff (s–1) and Kon = (1/τon − Koff)/[toxin] (nM–1S–1). Voltage-activation relationships were obtained by measuring steady-state Na+ currents elicited by step depolarizations from –110 to +80 mV with 10-mV increments. Peak conductance (GNa) was calculated from G = I/(V–Vrev), where I, V, and Vrev represent the current value, membrane potential, and reverse potential, respectively. The voltage of steady-state inactivation was estimated using a double-pulse protocol with currents elicited by a 20-millisecond depolarizing potential of 0 mV following a 500-millisecond prepulse to potentials from –130 to –10 mV using 10-mV increments. Then, voltage dependence of activation and inactivation were determined in the absence or presence of Tp1a (5-minute exposure) with the cells before application of the voltage protocols. Experimental data were analyzed using QPatch Assay Software version 5.0 (Sophion).

Cav1, Cav2, and Nicotinic Acetylcholine Receptor Assays.

Ca2+ responses were measured with FLIPRTETRA and Calcium 4 dye (Molecular Devices) using the neuroblastoma cell line SH-SY5Y as previously reported (Vetter and Lewis, 2010; Sousa et al., 2013). SH-SY5Y cells were plated at 40,000 cells per well in 384-well flat clear-bottom black plates (Corning) and cultured at 37°C in a humidified 5% CO2 incubator for 48 hours. The medium was removed and cells loaded with 20 μl/well Calcium 4 dye reconstituted in assay buffer containing (in millimolar) 140 NaCl, 11.5 glucose, 5.9 KCl, 1.4 MgCl2, 1.2 NaH2PO4, 5 NaHCO3, 1.8 CaCl2, and 10 HEPES (pH 7.4) and incubated for 30 minutes at 37°C in a humidified 5% CO2 incubator. For voltage-gated calcium channel 1 (CaV1) assays, 1 μM ω-conotoxin CVID (CaV2.2 blocker) was added to the dye, and for the CaV2.2 assay, 10 μM nifedipine (CaV1 blocker) was added to the dye. For assay of the α7 nicotinic acetylcholine receptor (nAChR), 10 μM PNU-120596 (N-(5-chloro-2,4-dimethoxyphenyl)-N′-(5-methyl-3-isoxazolyl)-urea; an α7 agonist) was added to the dye. Ca2+ fluorescence responses were recorded using excitation of 470–495 nm and emission of 515–575 nm for 10 seconds to set the baseline, then again 600 seconds after addition of 5 μM Tp1a and for a further 300 seconds after addition of 90 mM CaCl2 for CaV, 30 μM choline for the α7 nAChR, or 30 μM nicotine for assay of the α3β2/4 nAChR.

Pain Behavioral Assessment In Vivo.

Spontaneous pain was induced using OD1, a selective and potent activator of NaV1.6/NaV1.7 (Maertens et al., 2006). OD1 (300 nM) ± rGly-Tp1a (1 μM, 300 nM, 100 nM) in a volume of 40 μl was administered by shallow subcutaneous (intraplantar) injection into the left hindpaw under light isoflurane anesthesia. Immediately after injection, mice were placed individually into polyvinyl boxes (10 × 10 × 10 cm), and spontaneous pain was quantified by counting the number of paw lifts, licks, shakes, and flinches by a blinded observer over a 40-minute period in 5-minute intervals from video recordings.

Structure Determination of Tp1a.

Either the C-terminal amide or acid form of synthetic Tp1a (sTp1a; 0.2 mg) was dissolved in 0.5 ml of 90% H2O/10% D2O at pH ∼3.5. Two-dimensional (2D) homonuclear 1H NMR data were recorded over the temperature range 15–35°C at 600 MHz on a Bruker Avance spectrometer (Billerica, MA) equipped with a cryoprobe. Recorded data sets included 2D TOCSY (mixing time 80 milliseconds), 2D nuclear overhauser effect spectroscopy (mixing time 200 milliseconds), and 2D DQF-COSY. Data were recorded with 4K data points in the direct dimension and 512 increments in the indirect dimension over a sweep width of 12 ppm. All data were referenced to the water signal (4.768 ppm at 25°C).

For structure determination, interproton distances were derived from peak volumes in a nuclear overhauser effect spectroscopy spectrum using automated assignment strategies in the program CYANA (Guntert, 2004). Dihedral-angle restraints were derived from amide-proton coupling constants obtained from one-dimensional and DQF-COSY spectra. Hydrogen bonds were identified by analysis of amide-proton temperature coefficients. Amide protons with a temperature coefficient more than –4.6 ppb/K are likely to be involved in hydrogen bonds (Cierpicki and Otlewski, 2001); for such amides where suitable hydrogen bond acceptors could be identified in preliminary structure calculations, hydrogen-bond restraints were included (Wang et al., 2000). The final family of structures was calculated using the program CNS (Brünger et al., 1998) using protocols from the RECOORD database (Nederveen et al., 2005) and refinement in explicit water, as described previously (Conibear et al., 2012). From a family of 50 calculated structures, the 20 structures with the highest stereochemical quality as judged by MolProbity (Chen et al., 2010) were selected to represent the solution structure of Tp1a. Atomic coordinates and NMR chemical shifts have been submitted to the Protein Data Bank and Biological Magnetic Resonance Data Bank, under accession numbers PDB 2mxm and BMRB 25419, respectively.

Data Analysis.

Curve fitting was achieved using GraphPad Prism version 6 (GraphPad Software Inc., San Diego, CA) with nonlinear regression with log[inhibitor] versus normalized response and variable Hill slope for dose responses, Boltzmann sigmoidal equation for voltage dependence of activation and inactivation analysis, and exponential one-phase decay or association for on- and off-rate analysis. Data were represented as the mean ± S.E.M. from at least n = 3 replicates. For the in vivo experiments, data were presented as the mean ± S.E.M. and statistical significance determined by analysis of variance with Dunnett’s post test (n = 3–9).

Results

Screen for hNaV1.7 Inhibitors.

Venoms from 40 species of spiders that inhibited veratridine-evoked NaV channel activity (Klint et al., 2015) were rescreened in an improved assay in which hNaV1.7 was specifically activated using a combination of OD1 and veratridine. Using this approach, 18 spider venoms were found to potently inhibit hNaV1.7 (at least 90% inhibition) at concentrations of 250 and 25 ng/μl (unpublished data). The venom of T. pruriens potently inhibited hNaV1.7 (Fig. 1), consistent with the presence of the previously characterized hNaV1.7 inhibitors ProTx-I and ProTx-II (Middleton et al., 2002). Assay-guided fractionation was used to confirm whether these toxins accounted for the full activity of T. pruriens venom.

Fig. 1.
Fig. 1.

Screening of T. pruriens spider venom against hNaV1.7 endogenously expressed in SH-SY5Y cells. Effect of venom from T. pruriens on hNaV1.7 showing 100 and 90% inhibition at 250 (●) and 25 (+) ng/μl, respectively, and loss of activity at 2.5 ng/μl (★). The positive control for hNaV1.7 activation was veratridine plus OD1 (×).

Peptide Toxin Identification.

Fractionation of T. pruriens venom using C18 RP-HPLC revealed five major peaks (Fig. 2A), with three fractions inhibiting hNaV1.7 (Fig. 2B). The earliest eluting of these peptides, Tp1a, corresponded to the second dominant peak with a retention time of 26.9 minutes, whereas the more hydrophobic ProTx-I and ProTx-II eluted at 40.3 and 51.9 minutes, respectively. The reduced hydrophobicity of Tp1a encouraged further studies on this peptide due to its potentially more straightforward synthesis. Native Tp1a had a monoisotopic mass of 3799.73 Da as observed by matrix-assisted laser desorption/ionization–TOF and electrospray mass spectrometry (Fig. 2C). Edman degradation, tandem mass spectrometry, and amino acid analysis revealed that Tp1a is a 33-residue peptide with a calculated monoisotopic oxidized mass of 3799.70 Da, consistent with the presence of three disulfide bonds and an amidated C terminus (Fig. 3A).

Fig. 2.
Fig. 2.

Isolation of novel hNaV1.7 inhibitor from venom of the tarantula T. pruriens. (A) RP-HPLC fractionation of crude T. pruriens venom (1 mg) on a Phenomenex analytical C18 column using the gradient of solvent B indicated by the dashed line at a flow rate of 1 ml/min. Fractions were manually collected and screened for hNaV1.7 inhibition. Fractions found to inhibit hNaV1.7 are shaded dark gray. (B) Effect of RP-HPLC fractions on hNaV1.7 activity in SH-SY5Y cells as measured on a FLIPR. Fractions eluting at 26.9, 40.3, and 51.9 minutes potently inhibited hNaV1.7. The dominant masses found by matrix-assisted laser desorption/ionization–TOF analysis of each of the active fractions are indicated. Masses of 3984 and 3823 Da correspond to ProTx-I and ProTx-II, respectively. (C) Analytical RP-HPLC chromatogram and matrix-assisted laser desorption/ionization–TOF mass spectrum of RP-HPLC fraction 12 showing single peak and monoisotopic mass of 3799 Da that corresponds to Tp1a peptide.

Fig. 3.
Fig. 3.

Determination of sequence of Tp1a and comparison with other spider toxins. (A) Determination of the amino acid sequence of Tp1a. The sequence of the first 31 amino acids was obtained by Edman degradation. The last two amino acids were determined using a combination of tandem mass spectrometry (MS nanospray) and amino acid analysis (AAA). (B) Alignment of Tp1a with spider venom peptides having at least 40% sequence identity. Identical residues are shown in bold and cysteines are gray. The percent identity relative to Tp1a and activity reported for each peptide is shown on the far right. Peptide sequences were obtained from the ArachnoServer database (www.arachnoserver.org) (Herzig et al., 2011). Asterisks denote C-terminal amidation. (C) Alignment of the sequence of Tp1a with those of ProTx-I (β/ω-TRTX-Tp1a) and ProTx-II (β/ω-TRTX-Tp2a), two NaV channel inhibitors previously isolated from the same venom.

Tp1a belongs to NaSpTx family 1, a family of spider venom–derived NaV channel toxins that comprise 33–35 residues with three disulfide bonds that form a hyperstable inhibitor cystine knot (ICK) motif (Klint et al., 2012). The closest homologs of Tp1a are ω-TRTX-Gr2b (63% identity), β-TRTX-Ps1a (58% identity), and ω-TRTX-Gr2a (58% identity) (Fig. 3B). Surprisingly, Tp1a is very dissimilar to ProTx-I (25% identity) and ProTx-II (38% identity) from the same spider, which belong to NaSpTx families 2 and 3, respectively (Klint et al., 2012) (Fig. 3C).

Recombinant Expression and Chemical Synthesis of Tp1a.

Recombinant Tp1a was produced by periplasmic expression of a MBP-Tp1a fusion protein in E. coli. A non-native N-terminal Gly residue was added to facilitate TEV protease cleavage of the fusion protein since this is the preferred residue in the P1' position of the TEV protease recognition site. The MBP-Tp1a fusion protein was purified on a nickel column, then TEV cleavage was used to release unfused rGly-Tp1a (Fig. 4A). Only 50% of the fusion protein was cleaved, giving a yield of 0.4 mg/l rGly-Tp1a after final purification by RP-HPLC; the observed monoisotopic mass of 3858.23 Da is consistent with the presence of the non-native N-terminal Gly residue (Fig. 4B). rGly-Tp1a was produced as a single isomer that eluted on RP-HPLC ∼1 minute after native Tp1a (nTp1a). The recombinant peptide was lyophilized and stored at 4°C. Chemical synthesis was used to generate the C-terminal amide and acid forms of Tp1a, of which the amidated form coeluted with nTp1a (Fig. 5).

Fig. 4.
Fig. 4.

Production of recombinant Tp1a. (A) SDS-PAGE gel showing MBP-Tp1a fusion protein purified via nickel affinity chromatography (lane 1) and the efficiency of TEV protease cleavage of the fusion protein (∼50%) (lane 2). (B) RP-HPLC purification of cleaved recombinant rGly-Tp1a was performed using a Vydac 218TP C18 column with a two-step gradient of 5–50% solvent B over 45 minutes followed by 50–80% solvent B over 8 minutes. Matrix-assisted laser desorption/ionization–TOF mass spectrometry yielded a monoisotopic mass of 3858 Da, consistent with the calculated mass for rGly-Tp1a (3857.70).

Fig. 5.
Fig. 5.

Comparison of native and synthetic Tp1a. Analytical RP-HPLC chromatograms of native amidated Tp1a (dashed trace), synthetic Tp1a-NH2 (gray trace), and coelution of synthetic Tp1a-NH2 and native Tp1a (black trace). RP-HPLC was performed on a Shimadzu LC20AT system using a Thermo Hypersil GOLD C18 column (2.1 × 100 mm; Thermo Fisher Scientific, Waltham, MA) heated at 40°C. Peptides were eluted using a gradient of 5–50% B over 45 minutes with a flow rate of 0.3 ml/min.

Effect of Tp1a on hNaV1.7.

Tp1a activity was further investigated by whole-cell patch-clamp studies of CHO cells expressing hNaV1.7 (Fig. 6). Current traces in the presence and absence of nTp1a, rGly-Tp1a, sTp1a-OH, and sTp1a-NH2 are shown in Fig. 6A. Native, recombinant, and the synthetic C-terminal acid and amide forms of Tp1a inhibited hNaV1.7 with an IC50 of 2.1 ± 1.3 (n = 6), 9.5 ± 3.4 (n = 7), 11.5 ± 3.9 (n = 6), and 2.5 ± 0.8 (n = 6) nM, respectively (Fig. 6B).

Fig. 6.
Fig. 6.

Inhibition of hNaV1.7 by Tp1a. (A) Representative records of Na+ currents before (dashed traces) and after (black traces) addition of 2 nM native and synthetic Tp1a-NH2 or 10 nM recombinant and synthetic Tp1a-OH. Holding potential was –80 mV. (B) Concentration-response curves for inhibition of hNaV1.7 by native, recombinant, and synthetic acid and amidated Tp1a; the IC50 values calculated using I/Imax values and nonlinear regression were 2.1 ± 1.3 (n = 6), 9.5 ± 3.4 (n = 7), 11.5 ± 3.9 (n = 6), and 2.5 ± 0.8 (n = 6) nM (mean ± S.D.), respectively.

Effect of Tp1a on Activation and Inactivation of hNaV1.7.

Most NaSpTxs are so-called gating modifiers that modulate NaV channel activity by modifying the voltage dependence of activation and/or inactivation (Catterall et al., 2007). Figure 7 shows conductance-voltage and current-voltage relationships in the presence and absence of Tp1a. A Boltzmann sigmoidal fit showed a small shift in the voltage of activation and inactivation to more negative potentials in the presence of nTp1a (−8.89 and −7.24 mV, respectively) and sTp1a-NH2 (–5.1 mV) (Fig. 7, A and B). Similar changes in the activation and inactivation of hNaV1.7 were observed in the presence of sTp1a-OH, with ΔV1/2 shifted –5.27 and –6.52 mV, respectively (Fig. 7C). rGly-Tp1a had a more pronounced effect on hNaV1.7 inactivation, with ΔV1/2 shifted by –13.9 mV, and a lesser effect on activation, with ΔV1/2 shifted –3.3 mV (Fig. 7D).

Fig. 7.
Fig. 7.

Gating properties of Tp1a. Data (mean ± S.D., n = 3) for nTp1a (A), sTp1a-NH2 (B), sTp1a-OH (C), and rGly-Tp1a (D) are plotted as G/Gmax or I/Imax. Tp1a had no significant effect on the voltage dependence of steady-state activation or inactivation. Cells were held at –80 mV. Steady-state kinetics were estimated by currents elicited at 10-mV increments ranging from –110 to +80 mV. Conductance was calculated using G = I/(V–Vrev) in which I, V, and Vrev are the current value, membrane potential, and reverse potential, respectively. The voltage dependence of inactivation was estimated using a double-pulse protocol where currents were elicited by a 20-millisecond depolarizing potential of 0 mV following a 500-millisecond prepulse at potentials ranging from –130 to –10 mV with 10-mV increments. The ΔV1/2 for activation and inactivation were −8.89 and −7.24 mV for nTp1a (A), –5.1 and –5.1 mV for sTp1a-NH2 (B), –5.27 and –6.52 mV for sTp1a-OH (C), and –3.3 and –13.9 mV for rGly-Tp1a (D), respectively.

Kinetics of Tp1a Inhibition of hNaV1.7.

hNaV1.7 current inhibition and dissociation constant Kd were estimated following application and washout of Tp1a (Figs. 8 and 9; Table 1). The association rate (Kon) was ∼10-fold faster for nTp1a and sTp1a-NH2 compared with sTp1a-OH and rGly-Tp1a at nonsaturating concentrations (Fig. 8; Table 1). These findings reveal a critical role for the C-terminal functionality in determining the kinetics of Tp1a action on hNaV1.7. Inhibition of hNaV1.7 by recombinant and synthetic Tp1a-OH was not completely reversible, with sodium currents recovered to ∼60 and ∼45% of control levels, respectively (n = 3), after 70-minute washout, whereas inhibition by native and synthetic Tp1a-NH2 was quasi-irreversible (n = 3) (Fig. 9). The calculated Koff values were 3.11 ± 0.2 × 10−4 s–1 and 3.46 ± 0.5 × 10−4 s–1 for sTp1a-OH and rGly-Tp1a, respectively (Table 1).

Fig. 8.
Fig. 8.

On-rate of Tp1a inhibition of hNaV1.7. Measurement of on-rates for various forms of Tp1a. Na+ currents were recorded every 15 milliseconds soon after toxin addition. The calculated on-rates were 2.45, 1.63 × 103, and 2.01 × 103 minutes for nTp1a at 20, 2, and 0.2 nM, respectively (A); 2.28, 3.13 × 103, and 3.49 × 103 minutes for sTp1a-NH2 at 20, 2, and 0.2 nM, respectively (B); 3.15, 3.78 × 103, and 2.30 × 103 minutes for rGly-Tp1a at 150, 15, and 1.5 nM, respectively (C); and 2.40, 2.78 × 101, and 3.83 × 103 minutes for sTp1a-OH at 150, 15, and 1.5 nM, respectively (D).

Fig. 9.
Fig. 9.

Off-rate of Tp1a and analogs at hNaV1.7. Tp1a was applied at 20 nM for nTp1a and sTp1a-NH2 and 150 nM for rGly-Tp1a and sTp1a-OH and incubated for 10 minutes before Na+ currents were measured every 10 minutes during saline washout. sTp1a-OH and rGly-Tp1a bound reversibly with off-rates of 53.5 and 47.7 minutes, respectively, whereas nTp1a and sTp1a-NH2 showed quasi-irreversible binding to hNaV1.7.

View this table:
TABLE 1

Kinetics of inhibition of hNaV1.7 by nTp1a, sTp1a-NH2, sTp1a-OH, and rGly-Tp1a

Tp1a was applied at 0.2, 2, and 20 nM (nTp1a and sTp1a-NH2) or 1.5, 15, and 150 nM (sTp1a-OH and rGly-Tp1a), then sodium currents were measured. The kinetics of inhibition and recovery of inhibition were determined from the I/Imax as a function of time from traces shown in Fig. 8, fitted to a single exponential fit. Values are from at least three independent experiments (mean ± S.D.).

Subtype Selectivity of Tp1a.

The subtype selectivity of Tp1a was first examined by measuring changes in the membrane potential in HEK 293 cells expressing NaV α subunits coexpressed with the β1 subunit, except for NaV1.6, which was expressed in the absence of subunits (Table 2). rGly-Tp1a and sTp1a-OH had the highest potency for hNaV1.7 (IC50 1.1 ± 0.13 and 1.5 ± 0.4 μM, respectively), followed by hNaV1.2 = hNaV1.1 > hNaV1.6. No significant activity was observed against hNaV1.3, hNaV1.4, hNaV1.5, or hNaV1.8 at up to 5 μM rTp1a and sTp1a-OH. sTp1a-NH2 was more potent against all NaV subtypes, with the highest potency against hNaV1.7 (IC50 0.22 ± 0.05 μM) and hNaV1.6 (IC50 0.29 ± 0.05 μM) and less potency at hNaV1.2 > hNaV1.1 > hNaV1.3. No activity was observed at hNaV1.4, hNaV1.5, and hNaV1.8 at up to 5 μM sTp1a-NH2.

View this table:
TABLE 2

Tp1a activity on NaV channels evaluated by fluorescent imaging and electrophysiology represented as IC50 values

For fluorescent assays, hNaV recombinant cell lines were incubated with rGly-Tp1a, sTp1a-NH2, or sTp1a-OH for 30 minutes followed by activation with veratridine/OD1. Fluorescent intensity was normalized against positive and negative controls and maximum response from 5-minute reading after NaV activation used for plotting the concentration-response curves. For electrophysiology assays, NaV recombinant cell lines held at –80 mV were incubated with rGly-Tp1a, sTp1a-OH, or sTp1a- NH2 for 5 minutes followed by a prepulse of –120 mV for 200 milliseconds and activation with a single pulse of 0 mV for 20 milliseconds. I/Imax values were used for plotting the concentration-response curves. Values are from at least three independent experiments (mean ± S.D.).

Tp1a activity at other NaV channel subtypes was also investigated using whole-cell patch-clamp electrophysiology (Table 2). rGly-Tp1a inhibited hNaV1.1 with an IC50 of 59.8 ± 26 nM, whereas sTp1a-OH inhibited this subtype with an IC50 of 100.9 ± 30 nM. In contrast to the fluorescent assays, rGly-Tp1a and sTp1a-OH were shown to be active at hNaV1.3 by electrophysiology, with IC50 values of 21.9 ± 4.02 and 41.3 ± 4.7 nM, respectively. Tp1a-NH2 was 3.5- to 9-fold more potent than Tp1a-OH, with IC50 values of 11.3 ± 1.2 nM for hNaV1.1 and 11.5 ± 3.0 nM for hNaV1.3. No activity was observed at hNaV1.5 for either rGly-Tp1a or for the acid and amidated forms of sTp1a (in the presence of up to 500 nM). Activity at hNaV1.2, hNaV1.4, and hNaV1.8 was not investigated using electrophysiology.

Effect of Tp1a on Cav Channels and nAChRs.

We investigated the activity of Tp1a on CaV channels and nAChRs known to be involved in pain pathways and muscle contraction (Zamponi et al., 2009; Hurst et al., 2013). Tp1a did not affect CaV or nAChR activity at concentrations up to 5 μM (unpublished data). Currents from these channels were fully inhibited by 10 μM nifedipine for CaV1, 1 μM ω-conotoxin CVID for CaV2, and 10 μM tubocurarine for the α3β4/2 and α7 nAChRs.

Antinociceptive Effects of Tp1a in a Mouse Model of Pain.

Intraplantar injection of the NaV1.7 enhancer OD1 in mice led to rapid development of spontaneous pain as evidenced by flinching, lifting, licking, and shaking of the affected hindpaw, consistent with the role of NaV1.7 in pain. Intraplantar injection of rGly-Tp1a at 1 μM (40 pmol in a 40-μl injection) and 300 nM (12 pmol in a 40-μl injection) significantly reduced spontaneous pain behavior in a concentration-dependent manner (Fig. 10A; OD1, 103 ± 6 flinches/10 minutes; 1 μM Tp1a, 11 ± 3 flinches/10 minutes; 300 nM Tp1a, 49 ± 17 flinches/10 minutes, P < 0.05), and this reduction in pain behavior persisted for 25 minutes after injection of the highest concentration of Tp1a (Fig. 10B).

Fig. 10.
Fig. 10.

Antinociceptive effects of Tp1a. (A) Intraplantar injection of the NaV1.7 activator OD1 (300 nM) led to rapid development of nocifensive behavior in mice that slowly declined over 40 minutes. This spontaneous pain behavior, measured by the number of paw lifts, licks, shakes, and flinches, was significantly attenuated in a concentration-dependent manner by coadministration of Tp1a when compared with vehicle control. (B) The reduction in spontaneous pain behavior persisted for 25 minutes after injection of the highest concentration of Tp1a (1 μM). Data are presented as the mean ± S.E.M. of 3–9 mice/group. Statistical significance was determined by analysis of variance with Dunnett’s post test; *P < 0.05 compared with control.

3D Structure of Tp1a.

The structure of Tp1a was examined using 2D 1H NMR. Hα chemical shifts, which are sensitive to structural changes (Wishart et al., 1992), indicate that the amidated and acid forms of Tp1a have identical folds, with the only minor differences due to the presence or absence of the proximal C-terminal negative charge (Fig. 11A). Thus, the difference in pharmacology between these two forms of the peptide is not caused by structural differences.

Fig. 11.
Fig. 11.

3D structure of Tp1a. (A) Comparison of 1Hα chemical shifts for sTp1a-NH2 and sTp1a-OH, indicating that C-terminal amidation does not alter the 3D fold of Tp1a. (B) Ensemble of 20 structures chosen to represent the solution structure of Tp1a. Disulfide bonds are shown in yellow. (C) Schematic representation of Tp1a structure showing β-strands (magenta) and disulfide bonds (yellow). (D) Surface representation of the structures of Tp1a (left), μ-TRTX-Hhn1b (center), and μ-TRTX-Hs2a (right). Positively and negatively charged residues are highlighted in blue and red, respectively. Residues shown to be critical for inhibition of NaV1.7 include K27 and R29 for μ-TRTX-Hhn1b (Li et al., 2004) and K18, R26, and K32 for μ-TRTX-Hs2a (Minassian et al., 2013; Revell et al., 2013).

The 3D structure of the acid form of Tp1a was determined using NMR. The final ensemble of 20 structures is shown in Fig. 11, B and C, and structural statistics are presented in Table 3. The structure is in excellent agreement with the experimental restraints, and it has good covalent geometry.

View this table:
TABLE 3

Structural statistics for the ensemble of Tp1a structures

Based on sequence homology, we anticipated that Tp1a would adopt an ICK fold. This was confirmed by NMR analysis, which revealed that the peptide contains an ICK motif in which the Cys2-Cys17 and Cys9-Cys22 disulfide bonds form a 14-residue ring that is bisected by the Cys16-Cys29 disulfide bond. Elements of secondary structure include a β-sheet comprising residues 20–23 and 29–31, and several β-turns comprising residues 4–7 and 17–20 (type II) and residues 10–13 and 23–26 (type I). The latter turn is present in the least well defined region of the structure, and based on the significant line broadening observed for His26, it appears that this region is dynamic in solution. In addition to the three disulfide bonds, the structure is stabilized by a large number of hydrogen bonds, including both backbone and side-chain groups.

We compared the molecular surface of Tp1a to that of μ-TRTX-Hhn1b and μ-TRTX-Hs2a (Fig. 11D). As for Tp1a, μ-TRTX-Hhn1b and μ-TRTX-Hs2a preferentially inhibit NaV1.2, NaV1.3, and NaV1.7, with no effect on NaV1.4 and NaV1.5 (Xiao et al., 2008; Cai et al., 2015). Mutations of μ-TRTX-Hhn1b revealed that Lys27 and Arg29 are critical for activity on NaV channels (Li et al., 2004), whereas an alanine scan of μ-TRTX-Hs2a revealed that residues Lys18, Arg26, Lys32, and Trp30 are critical for NaV channel inhibition (Minassian et al., 2013; Revell et al., 2013). Equivalent residues in the Tp1a structure are Lys8, Arg12, and Lys30, which we propose are likely to be important for Tp1a inhibition of hNaV1.7.

Discussion

NaV channels have been extensively studied using neurotoxins from spiders, cone snails, centipedes, and scorpions that have high affinity and specificity for this class of ion channels (King, 2011). By using an in-house high-throughput screen, we isolated a novel spider venom peptide (Tp1a) that potently inhibited hNaV1.7 and reversed pain responses in vivo.

Isolation and Production of Tp1a.

Tp1a was identified in the venom of the tarantula T. pruriens, from which the NaV channel inhibitors ProTx-I and ProTx-II were previously identified (Middleton et al., 2002). However, despite their common origin, Tp1a has a much higher sequence identity with the NaV channel inhibitors ω-TRTX-Gr2b, β-TRTX-Ps1a, and ω-TRTX-Gr2a (Fig. 3) than with ProTx-I and ProTx-II. Thus, the venom of T. pruriens contains Tp1a from NaSpTx family 1, ProTx-I from NaSpTx family 2, and ProTx-II from NaSpTx family 3, suggesting that NaV channel inhibition plays a key role in immobilization of prey by this spider.

Recombinant Tp1a was readily produced using a bacterial expression system as reported for other ICK peptides (Saez et al., 2011; Klint et al., 2013), although the recombinant version had a non-native N-terminal Gly residue, and the C terminus was not amidated as in the native toxin. In contrast, chemical synthesis of Tp1a allowed production of both the native amidated form as well as the corresponding free carboxyl form without the extra Gly residue. Cell-based FLIPR assays revealed that both recombinant and synthetic carboxylated Tp1a preferentially inhibited hNaV1.7, whereas the more potent amidated form preferentially inhibited hNaV1.7 and hNaV1.6 (Table 2). C-terminal amidation also significantly enhanced potency at hNaV1.3 compared with the acid forms of Tp1a.

Pharmacology of Tp1a over NaV Channels.

Whole-cell patch-clamp electrophysiology was used to further characterize the subtype selectivity of Tp1a. Once again, the amidated form was found to be more potent (Fig. 6). Indeed, the potency of amidated Tp1a (IC50 2.1 and 2.5 nM for native and synthetic Tp1a, respectively) is comparable or better than the most potent hNaV1.7 inhibitors described to date, including ProTx-I (IC50 51 nM), ProTx-II (IC50 0.3 nM), and huwentoxin-IV (IC50 26 nM) as measured using similar approaches (Middleton et al., 2002; Xiao et al., 2008). Curiously, Tp1a induced no significant changes in the voltage dependence of activation or steady-state inactivation of hNaV1.7 (Fig. 7). This suggests that Tp1a may act like huwentoxin-IV, which binds to the neurotoxin site 4 located at the extracellular S3-S4 loop of domain II to trap the voltage sensor in a closed configuration (Xiao et al., 2008).

C-terminal amidation increased Kon and slowed Koff for Tp1a, such that inhibition of hNaV1.7 by amidated Tp1a was quasi-irreversible (Figs. 8 and 9; Table 1). These kinetic data, the first to be reported for NaSpTx family 1 toxins, support the enhanced potency observed for amidated Tp1a. Amidation of Tp1a enhanced affinity at all targeted subtypes, whereas the carboxyl form suffered its most dramatic loss of affinity at hNaV1.6 and hNaV1.3. These shifts in potency were observed in both electrophysiology and fluorescent imaging assays, despite the significant differences in the IC50 values obtained with the approaches. Such differences in electrophysiology and fluorescent imaging assays for ion channels have been reported previously (Mathes et al., 2009; Terstappen et al., 2010), but the underlying cause remains to be elucidated.

Structure of Tp1a Reveals Typical ICK Motif.

The ICK scaffold is the fold adopted by the vast majority of spider venom peptides (King and Hardy, 2013). The three canonical disulfide bonds in the ICK motif form a knot in which a ring formed by the C1-C4 and C2-C5 disulfide bonds and the intervening sections of polypeptide backbone is penetrated by the third disulfide bridge (C3-C6). ICK peptides differ in the size of the intercysteine loops and the length of the N- and C-terminal regions, but the ICK motif is expected to provide most of these peptides with high levels of chemical, thermal, and biologic stability (Saez et al., 2011). As for other members of NaSpTx family 1, such as hainantoxin-IV (μ-TRTX-Hhn1b) and huwentoxin-IV (μ-TRTX-Hs2a) (Peng et al., 2002), we found that Tp1a also contains an ICK motif. The β-hairpin loop (i.e., intercystine loop 4) of NaSpTx family 1 toxins has been identified as an important region for interaction with NaV channels, and it will be interesting to see whether this is the case for Tp1a. The successful recombinant expression and chemical synthesis of Tp1a demonstrated here will facilitate future studies of structure-activity relationships by enabling rapid and affordable production of Tp1a analogs. Alterations in the C-terminal region can also influence the potency and selectivity of these toxins. These alterations can cause significant structural modifications as reported for huwentoxin-IV, or not affect the structure and still generate striking pharmacological changes (Minassian et al., 2013; Revell et al., 2013). We found that the C terminus of Tp1a is functionally critical, with C-terminal amidation increasing the potency of the toxin and reducing its reversibility.

Analgesic Potential of Tp1a.

Given the important role identified for NaV1.7 in nociception, we tested the analgesic efficacy of Tp1a in a rodent pain model in which nocifensive behavior (paw lifting, licking, and flinching) is elicited by intraplantar administration of the NaV1.7 activator OD1. Intraplantar administration of Tp1a reduced the nocifensive behavior induced by OD1 by ∼90%, and this effect persisted up to 25 minutes after Tp1a administration. This suggests that Tp1a can effectively inhibit NaV1.7 at peripheral sensory nerve endings in the skin. Nonetheless, future detailed assessment of the analgesic potential of Tp1a in more sophisticated disease-specific pain models will provide additional insight into the clinical potential of this peptide for the treatment of chronic pain.

Conclusions.

We used a novel high-throughput screening strategy to rapidly isolate a potent NaV1.7 inhibitor from spider venom. The pharmacological properties of Tp1a evaluated by fluorescent imaging and electrophysiology approaches revealed a crucial role of C-terminal amidation in potency, affinity, and NaV subtype selectivity of spider toxins. Tp1a was analgesic in a NaV1.7-specific model of pain, and its well defined structure may help guide the development of improved NaV1.7 inhibitors.

Acknowledgments

The authors thank members of the Deutsche Arachnologische Gesellschaft for providing spiders, especially Ingo Wendt (Schorndorf, Germany) for T. pruriens. The authors thank Dr. Alun Jones for help with mass spectrometry, and Niraj Bende (Institute for Molecular Bioscience, The University of Queensland, Brisbane, Australia) and Richard Allen (Australian National University, Canberra, Australia) for fractionation of T. pruriens venom. The authors thank members of the Australian Proteome Analysis Facility. Access to the Australian Proteome Analysis Facility is facilitated by support from the Australian Government's National Collaborative Research Infrastructure Strategy.

Authorship Contributions

Participated in research design: Cardoso, Lewis, King.

Conducted experiments: Cardoso, Dekan, Rosengren, Erickson, Vetter, Deuis.

Contributed new reagents or analytic tools: Herzig.

Wrote or contributed to the writing of the manuscript: Cardoso, Lewis, King, Alewood.

Footnotes

    • Received February 2, 2015.
    • Accepted May 15, 2015.

  • This work was supported by the Australian Research Council Discovery Project [Grant DP110103129], Linkage Project [Grant LP130101143], Future Fellowships to K.J.R. and I.V., and the Australian National Health and Medical Research Council Principal Research Fellowships to R.J.L., G.F.K., and P.A. The Australian Proteome Analysis Facility is supported under the Australian Government’s National Collaborative Research Infrastructure Strategy.

  • dx.doi.org/10.1124/mol.115.098178.

Abbreviations

ACN
acetonitrile
calc. (avg)
calculated (average)
CHO
Chinese hamster ovary
2D
two-dimensional
3D
three-dimensional
ESI
electrospray ionization
FBS
fetal bovine serum
FLIPR
fluorescent imaging plate reader
HEK 293
human embryonic kidney 293
HPLC
high-performance liquid chromatography
ICK
inhibitor cystine knot
MBP
maltose binding protein
MS
mass spectrometry
nAChR
nicotinic acetylcholine receptor
NaSpTx
NaV channel toxin
PNU-120596
N-(5-chloro-2,4-dimethoxyphenyl)-N′-(5-methyl-3-isoxazolyl)-urea
RP
reversed phase
TEV
tobacco etch virus
TFA
trifluoroacetic acid
TN
tris NaCl
TOF
time of flight
Trt
triphenylmethyl

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New Sodium Channel Inhibitor from Thrixopelma pruriens

Fernanda C. Cardoso, Zoltan Dekan, K. Johan Rosengren, Andelain Erickson, Irina Vetter, Jennifer R. Deuis, Volker Herzig, Paul F. Alewood, Glenn F. King and Richard J. Lewis
Molecular Pharmacology August 1, 2015, 88 (2) 291-303; DOI: https://doi.org/10.1124/mol.115.098178
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