Chemoselective tarantula toxins report voltage activation of wild-type ion channels in live cells

Edited* by Richard W. Aldrich, The University of Texas at Austin, Austin, TX, and approved September 23, 2014 (received for review May 27, 2014)
October 20, 2014
111 (44) E4789-E4796


Electrically excitable cells, such as neurons, exhibit tremendous variation in their patterns of electrical signals. These variations arise from the collection of ion channels present in any specific cell, but understanding which ion channels are at the root of particular electrical signals remains a significant challenge. Here, we describe novel probes, derived from a tarantula venom peptide, that are able to report the activity of voltage-gated ion channels in living cells. This technology uses state-selective binding to optically monitor the activation of ion channels during cellular electrical signaling. Activity-reporting probes based on these prototypes could potentially identify when endogenous ion channels contribute to electrical signaling, thus facilitating the identification of ion channel targets for therapeutic drug intervention.


Electrically excitable cells, such as neurons, exhibit tremendous diversity in their firing patterns, a consequence of the complex collection of ion channels present in any specific cell. Although numerous methods are capable of measuring cellular electrical signals, understanding which types of ion channels give rise to these signals remains a significant challenge. Here, we describe exogenous probes which use a novel mechanism to report activity of voltage-gated channels. We have synthesized chemoselective derivatives of the tarantula toxin guangxitoxin-1E (GxTX), an inhibitory cystine knot peptide that binds selectively to Kv2-type voltage gated potassium channels. We find that voltage activation of Kv2.1 channels triggers GxTX dissociation, and thus GxTX binding dynamically marks Kv2 activation. We identify GxTX residues that can be replaced by thiol- or alkyne-bearing amino acids, without disrupting toxin folding or activity, and chemoselectively ligate fluorophores or affinity probes to these sites. We find that GxTX–fluorophore conjugates colocalize with Kv2.1 clusters in live cells and are released from channels activated by voltage stimuli. Kv2.1 activation can be detected with concentrations of probe that have a trivial impact on cellular currents. Chemoselective GxTX mutants conjugated to dendrimeric beads likewise bind live cells expressing Kv2.1, and the beads are released by channel activation. These optical sensors of conformational change are prototype probes that can indicate when ion channels contribute to electrical signaling.
Electrical signals traveling along excitable cell membranes activate voltage-gated ion channels (VGICs), which open their pores to create electrical feedback and trigger signaling cascades that lead to neurotransmitter release, hormone secretion, gene transcription, and other cellular responses. Many maladies result from aberrant VGIC signaling, including epilepsies, cardiac arrhythmias, and pain syndromes (1, 2). VGIC complements vary with cell type (3), making cell-specific channels important targets to selectively modulate pathophysiological electrical signals (4). There are over 80 VGIC pore-forming subunits encoded in the human genome (5), and many of these are coexpressed within the same cell (3), making it difficult to dissect specific channel contributions to electrical signaling. In neurons, the precise trafficking of VGIC subtypes to distinct cell regions such as the axon hillock, presynaptic terminals, and distal dendrites allows specific subtypes to control voltage changes in subcellular compartments (69). However, the presence of an ion channel subunit in a cell does not indicate whether it is activated under physiological conditions (1013). Determining which ion channels activate under particular physiological and pathophysiological conditions is a formidable challenge, yet this understanding is crucial for identification of channel subtypes as therapeutic targets to correct pathologies (4). More than three decades after the first ion channel was cloned, voltage clamp remains the primary method to determine which ion channels are activated by electrical stimuli, making assignment of electrical activity to specific channel subtypes a difficult and imprecise task. New tools to optically report activity of VGICs are needed to advance study of their contributions to electrical signaling.
We hypothesized that ion channel activity could be reported by imaging ligands whose affinity for channels is state-dependent. The venoms of many predatory creatures contain voltage sensor-targeting toxins (VSTs) that act by binding selectively to specific channel conformations (1421). Even at doses too low to have a significant physiological effect, a state-dependent VST will bind preferentially to a specific ion channel conformation. Thus, the propensity of a state-dependent VST to bind an ion channel is dependent on the channel's activation status, and tracking VST localization could reveal spatiotemporal patterns of channel activity.
An intensively studied class of VSTs is from spider venoms. Spider VSTs have an “inhibitory cystine knot” fold that is stabilized by disulfide bridges. These structurally similar VSTs act upon Ca2+, Na+, and K+ channels, some with remarkable specificity for particular channel subtypes (22). VSTs are water-soluble peptides that partition into the outer leaflet of the plasma membrane, where they bind to the extracellular edge of VGIC transmembrane segments that form voltage sensors (21, 2326). This functional requirement to be soluble in water and partition into membranes suggests a delicate balance between a lipophilic face to interact with membrane lipids and bind the channel's transmembrane segment, and a hydrophilic face essential to prevent toxin aggregation and retain solubility (Fig. 1A) (27, 28). This complex biochemistry has made VSTs refractory to labeling attempts, as chemical perturbations are likely to disrupt peptide folding or activity. However, if these chemistry challenges are overcome, the resultant tools could be valuable probes of ion channel activity.
Fig. 1.
Tarantula toxin retains bioactivity following chemoselective conjugation. (A) Schematic of a VST portrays partitioning into a cell membrane (dashed line). Star represents a reporter conjugate. (B) GxTX saturably shifts the Kv2.1 conductance–voltage relation. Whole-cell voltage-clamp recordings from Kv2.1. Vehicle (black circles), 100 nM GxTX (red circles), 1 µM GxTX (red triangles), n = 4. Lines are fits of Eq. S1. Vehicle Vmid = 8 ± 3 mV, z = 1.2 ± 0.1e+; 100 nM GxTX Vmid = 81.5 ± 0.4 mV, z = 1.21 ± 0.03e+; 1 µM GxTX Vmid = 72 ± 1 mV, z = 1.24 ± 0.05e+. Values normalized to maximum conductance in vehicle. (C) Sequence of GxTX. Bold indicates residues replaced by Cys(Acm). Black indicates an intracystine loop or terminus where at least one residue could be replaced with Cys(Acm) yet retain activity against Kv2.1 at 100 nM. Yellow is cystine. (D) Red ribbon, backbone trace of GxTX (red ribbon); sticks, side chains replaced by Cys(Acm). Colored as in C. (E) Surface mesh representation of GxTX. Side chains colored as in C. (F) Extracellular view of Rosetta model of GxTX (red) bound to S3b helix of a Kv2.1 channel (gray). (G) Rendering of GxTX-TMR (red) binding to voltage sensor paddle of Kv2.1 (gray). Dashed line represents external membrane boundary. (H) GxTX-PEG5K retains activity against Kv2.1. Vehicle (black): Vmid = 10 ± 1 mV, z = 1.17 ± 0.04e+; 1 µM GxTX-PEG5K (red): Vmid = 64 ± 2 mV, z = 1.4 ± 0.3e+; n = 5. (I) GxTX-dy550 retains activity against Kv2.1. Vehicle (black), 20 nM GxTX-dy550 (red), n = 4. Lines are fits of Eq. 1. Vehicle Vmid = 3.7 ± 0.4 mV, z = 1.72 ± 0.02e+; 20 nM GxTX-dy550 Vmid = 55 ± 1 mV, z = 1.04 ± 0.05e+.
We sought to develop a general VST labeling strategy by taking advantage of recent advances in bioconjugation chemistry, which have produced a series of novel chemoselective reactions for modification of peptides. Here, we demonstrate that a cystine knot VST can be modified with fluorophores and affinity probes in a two-step process involving the incorporation of a small chemoselective side chain, which allows the peptide to retain its activity and permits the attachment of reporter molecules. We find that these conjugates report the localization of Kv2 channels, and dynamically detect activation of these ion channels in living cells, suggesting an optical means to monitor activation of specific channel subtypes during neuronal signaling.


VSTs Retain Bioactivity After Chemoselective Modification.

We chose to synthesize VST reporters from the tarantula peptide guangxitoxin-1E (GxTX), a potent modulator that selectively binds Kv2 channels (29, 30). In combination with patch-clamp electrophysiology, GxTX pharmacology is currently the most stringent test of whether Kv2 channels contribute to electrical signaling (13). In neurons and pancreatic islet cells, GxTX inhibits delayed rectifier current with minimal effects on other current types and has been recently used to reveal unexpected contributions of Kv2 channels to electrical signaling (13, 31, 32). Based on the effects of GxTX on Kv2 channel gating and the behavior of radiolabeled GxTX (30), we suspected GxTX might bind much more weakly when the channels become activated by voltage. To test this, we synthesized fluorescently labeled GxTX to determine if this VST could optically report Kv2 channel localization and activation.
In an initial attempt at generating GxTX variants, we expressed GxTX with an N-terminal hexahistidine tag and a C-terminal protease cleavage site in bacteria (SI Materials and Methods), but this peptide had little activity against Kv2.1 channels (Fig. S1A). This suggested that attachment of reporters to side chains, rather than the termini, was most likely to result in bioactive GxTX. We turned to chemical synthesis of GxTX to exploit nonnative amino acids that could serve as chemoselective conjugation points. We synthesized a GxTX variant with its sole methionine replaced by an oxidation-resistant norleucine isostere and found it altered Kv2.1 function by producing a strikingly positive shift in activation voltage (Fig. 1B), similar to wild-type GxTX (27, 29, 33). To avoid methionine oxidation complications, norleucine 35 GxTX was used as the background for further variants. To identify GxTX residues that can be substantially modified without disrupting toxin folding or activity, we introduced chemoselective amino acids into a series of variants. Mutagenesis studies of homologous VSTs (28, 34) guided selection of residues on the hydrophilic face of GxTX (33) that might have minimal impacts on channel binding. We substituted orthogonally protected cysteine residues [Cys(Acm)] at seven hydrophilic sites (Fig. 1C) and tested for activity against Kv2.1 channels. Of these seven Cys(Acm) mutants, five acted similar to GxTX at 100-nM concentration, shifting the voltage midpoint of Kv2.1 conductance to greater than +40 mV (Fig. S1A). When Lys27 was replaced with Cys(Acm), more than 300 nM was required to induce a perceptible gating shift; GxTX with Asn32 replaced by Cys(Acm) was inactive at concentrations up to 1 μM. Regions of GxTX that were tolerant to perturbations were suggested by mapping substitutions onto a GxTX NMR structure (33) (Fig. 1 D and E).
We generated a speculative model of GxTX bound to its Kv2.1 channel binding site (35) using Rosetta computational methods (3639). The lowest energy resultant docked structure served as a structural hypothesis for how GxTX interacts with each subunit of Kv2.1 (Fig. 1F). Although GxTX stabilizes resting conformations of the voltage sensor (35), this model represents GxTX binding to what is proposed to be an activated conformation of the voltage sensor (40). If the orientation of the toxin toward the channel remains similar in different states, it could be useful for predicting points of attachment for reporter molecules. This docking configuration places intracystine loop 2 most distal from the channel and membrane, suggesting that bulky reporter conjugates attached to loop 2 might be less likely to interfere with channel binding (Fig. 1G).
To create probe attachment sites on loop 2, we incorporated chemoselective groups, either an alkyne or a free thiol, for conjugation of fluorescent and affinity probes. To add an alkyne, we synthesized GxTX with the unnatural amino acid propargylglycine, for labeling with azide-bearing probes using Cu(I)-catalyzed Click reactions. Whereas many Click variations have been reported, Cu(I) salts are known to catalyze the disproportionation of thiol compounds (41), and we sought to identify conditions that would neither reduce nor scramble the cystine knot disulfides. We found that addition of the water-soluble Cu(I)-chelator bis[(tertbutyltriazoyl)methyl]-[(2-carboxymethyltriazoyl)methyl]-amine (BTTAA) (42) yielded GxTX Click conjugates that retained function against Kv2.1 channels. We attached a monofunctional 5-kDa polyethylene glycol (PEG5K) by its azide group to GxTX Pra13 (Fig. S1 B and D). The product (GxTX-PEG5K) was active against Kv2.1 channels, thereby inhibiting channel opening by shifting the midpoint of conductance to voltages similar to wild-type GxTX (Fig. 1H).
As an alternate strategy for chemoselective conjugation, we revealed a free “spinster” thiol on the surface of the peptide (43). Removal of the Acm protecting group has typically been catalyzed by Ag+ and facilitated with excess reductant (44), or oxidant (45), but these conditions are incompatible with preserving existing disulfides while obtaining a reduced surface thiol. To achieve the required mixed redox state, we removed Acm after GxTX folding and disulfide formation by displacing the Acm group with Ag+ followed by precipitation of Ag+ with Cl under acidic conditions, breaking the Ag–S bond without reducing or scrambling the disulfides (Fig. S1C). Maleimide-functionalized fluorophores, including tetramethylrhodamine and dylight550, were condensed with the free thiol, and Cys13-fluorophore GxTX conjugates retained activity against Kv2.1 (Fig. 1I). These results demonstrate that variants of GxTX can be folded and chemoselectively ligated to reporter molecules while retaining bioactivity against ion channels.

Fluorescent Toxin Colocalizes with Kv2.1 Channels in Live Cells.

To determine whether these VST conjugates could identify ion channels in live cells, we tested whether GxTX–fluorophores would colocalize specifically with Kv2.1 ion channels. When two mammalian CHO-K1 cell lines, one expressing Kv2.1 and the other expressing a blue fluorescent protein (BFP), were briefly treated with 100 nM of a GxTX–fluorophore conjugate, the GxTX fluorescence remained associated only with Kv2.1-expressing cells (Fig. 2A). When CHO-K1 cells were transiently transfected with Kv2.1-GFP, the channels appeared in clusters on the cell surface (Fig. 2B). GxTX localized tightly with these clusters (Pearson’s overlap coefficient r = 0.796), whereas surrounding untransfected cells had little GxTX fluorescence. In living neurons, a similar phenomenon was seen (Fig. 2C), although the colocalization with Kv2.1-GFP was not as complete (Pearson’s overlap coefficient r = 0.589), possibly owing to the presence of endogenous Kv channels. Whether tarantula VSTs such as GxTX have a high affinity for channels is a point of active debate (27, 46, 47). GxTX, like many other tarantula peptides, partitions into the outer leaflet of the plasma membrane, and it has been difficult to discern how much of its affinity for channels is due to membrane partitioning versus channel binding interactions (27). The colocalization of GxTX with Kv2.1 in cultured cells and neurons demonstrates that a strong, specific interaction occurs between the tarantula peptide and the ion channel voltage sensor. This precise colocalization makes GxTX–fluorophores viable ion channel probes.
Fig. 2.
Fluorescent tarantula toxin reveals Kv localization in live cells. (A) GxTX binds selectively to cells expressing Kv2.1. Images of two cocultured CHO-K1 cell lines that express either Kv2.1 at the cell surface or BFP in the nucleus. (Top) Differential interference contrast image of a mixed culture. (Middle) Fluorescence from GxTX-dy550. (Bottom) Merge of GxTX-dy550 (red), with BFP fluorescence (blue). (B) GxTX labels Kv2.1 clusters. Confocal slice through confluent CHO cells, one of which is expressing EGFP-Kv2.1. (Top) EGFP-Kv2.1 (green). (Middle) GxTX-TMR (red). (Bottom) Merge. Pearson’s overlap coefficient r = 0.80. (C) GxTX labels Kv subunits on cultured hippocampal pyramidal neurons. (Top) EGFP-Kv2.1 (green). (Middle) GxTX-dy550 (red). (Bottom) Merge. Pearson’s overlap coefficient r = 0.59.

Toxin Binding is Dependent on Channel Activation.

We hypothesized that GxTX binding is coupled to channel activity: when the membrane voltage is depolarized, voltage sensors adopt activated conformations and the binding of the inhibitory VST is weakened (Fig. 3A). Studies have concluded that the affinity of ion channels for certain VSTs is voltage dependent and that depolarizing pulses can release inhibitory VSTs from the channels (1821), but the precise changes in affinity have never been measured. To measure the degree to which GxTX affinity depends on voltage activation, the fraction of Kv2.1 channels bound by GxTX was deduced from K+ current at different holding potentials (SI Materials and Methods). At a holding potential of −100 mV, GxTX was found to inhibit K+ current during brief test pulses with a half-maximal inhibitory concentration (IC50) of about 2 nM (Fig. 3B, black). In cells held at 0 mV, the IC50 rose beyond 100 nM (Fig. 3B, blue). This shift of GxTX affinity with voltage indicates that GxTX indeed binds to Kv2.1 in a conformation-dependent manner.
Fig. 3.
Affinity of toxin for voltage sensors is conformation dependent. (A) Schematic of GxTX portrays the variable affinity for the resting and activated conformations of Kv2.1. Upon depolarization, active channels (blue) bind GxTX (red) with a lower affinity. (B) Normalized dose–response profile of GxTX with Kv2.1 held at −100 mV (black) or 0 mV (blue). Lines are fit of Eq. S2 (solid lines) ± SEM (dotted lines). Kd (−100 mV) = 12.7 ± 0.9 nM and Kd (0 mV) = 826 ± 139 nM. *P = 0.0003 at 100 nM. (C) Representative association of 100 nM GxTX to cells held at −100 mV (black) or 0 mV (blue). Red bar indicates perfusion of 100 nM GxTX. Lines are fits of Eq. S3 where kon −100 mV = 3.13 × 105 ± 6.58 × 103 M-1⋅s−1 and kon 0 mV = 4.40 × 104 ± 1.81 × 103 M-1⋅s−1. (D) Association rates of GxTX at −100 mV (black) and 0 mV (blue) determined by Eq. S3. kon 100 mV = 3.34 × 105 ± 1.13 M-1⋅s−1 and kon 0 mV = 2.53 × 104 ± 1.18 M-1⋅s−1 were determined by a linear fit (solid line) ± SEM (dotted lines). *P = 0.005 at 100 nM. (E) Representative GxTX dissociation from cells held at −100 mV and given a 100-ms 0-mV test pulse every 10 s (black) or given a 5-s 0-mV step every 10 s (blue). Red bar indicates GxTX perfusion. Lines are fits of Eq. S3 where koff −100 mV = 2.94 × 10−3 ± 2.13 × 10−5⋅s−1 and koff 0 mV = 2.23 × 10−2 ± 6.16 × 10−4⋅s−1. (F) Summary of dissociation rates of GxTX at −100 mV (black) and with 0-mV pulses (blue) determined by Eq. S3. Mean (solid line) ± SEM (dotted lines). *P = 0.005.
To better understand the thermodynamics of GxTX interactions with Kv2.1, we further analyzed the dose dependence of electrophysiological responses. Whereas four tarantula VSTs may bind a Kv channel tetramer (Fig. 1F) (48), only one toxin is required to eliminate current at neutral voltage (27). Due to this 4:1 stoichiometry, conductance at 0 mV is related to the GxTX dissociation constant (Kd) by the fourth power of the probability of a subunit being bound (SI Materials and Methods, Eq. S2). When this established analysis (48) is applied to our dose–response data set, it reports a 65-fold shift in the Kd for GxTX at −100 mV compared with 0-mV holding potential (Fig. 3B, curves). Voltage activation by depolarization to 0 mV induces Kv2.1 channels to reach a new distribution between closed, open, and inactivated conformations. Toxin-treated channels do not appreciably open at 0 mV (Fig. 1B), suggesting the toxin binding will be even weaker to more activated conformations at positive voltages. Whereas the relative affinities of the many closed, open, and inactivated channel conformations could not be distinguished with these experiments, our results clearly indicate that voltage activation lowers the affinity of GxTX for Kv2.1.
To determine whether the voltage-dependent shift in the GxTX Kd was due to a change in the binding rate, dissociation rate, or both, we measured kinetics of Kv2.1 inhibition and recovery at different voltages, and calculated microscopic binding–dissociation rates. The time course of inhibition and recovery were fit with a function that extracts a microscopic binding rate (kon) and dissociation rate (koff) (SI Materials and Methods, Eq. S3). When cells were held at −100 mV, GxTX inhibition occurred faster than at 0 mV (Fig. 3C), consistent with the toxin's weaker affinity at this depolarized voltage. This suggests that access of the VST to its binding site is compromised when the voltage sensors are activated. At both voltages, the time course of inhibition accelerated as GxTX concentration increased (Fig. 3D). Association rate rose linearly with GxTX concentration, consistent with its expected first-order dependence (Fig. 3D). kon became 13-fold slower when the voltage was raised to 0 mV (Table 1). To determine koff accurately, dissociation was measured after toxin washout. The return of Kv2.1 current after GxTX washout was accelerated when long pulses to 0 mV were included in the voltage stimulus protocol (Fig. 3E; SI Materials and Methods). The calculated koff during these 0-mV pulses was 70-fold faster than at −100 mV (Fig. 3F). Thus, toxin binding and dissociation are both affected by voltage, which indicates that GxTX's state specificity is due to its slow binding and rapid dissociation from voltage-activated channels. Fundamentally, the ratio of koff/kon determines the Kd of a bimolecular process. At both voltages tested, the ratio of koff/kon (calculated from kinetics, Eq. S3) is within an order of magnitude of Kd (calculated from equilibrium measurements, Eq. S2), but the values do not precisely match. This difference suggests the thermodynamic model that forms the basis of parameter extraction requires further refinement. Our calculations assume that toxin binds to each of four channel subunits independently and that toxin affinity is static over time. The independence of a tarantula VST binding to Kv2.1's four subunits has been experimentally validated (16). However, the voltage-dependent dissociation of tarantula VSTs from mutant Kv2.1 channels was found to be dependent on pulse duration and frequency (21). Progression through multiple resting, activated, and inactivated conformations is expected to result in affinity change over time. Although our model is oversimplified, it captures the basic features of activation-driven affinity change. The parameters extracted from kinetic measurements with this model show that channel activation alters the rates of association and dissociation in rough agreement with equilibrium measurements, demonstrating that the toxin has high affinity for resting channels and low affinity for activated channels.
Table 1.
Experimental and calculated GxTX-norleucine 35 affinity for rKv2.1 channels heterologously expressed in CHO cells
Parameter−100 mV0 mV
Kd (nM)12.7 ± 0.9826 ± 139
koff (s−1)0.84 ± 0.18 × 10−338 ± 9 × 10−3
kon (M-1⋅s−1)330 ± 110 × 10325 ± 12 × 103
koff/kon (nM)2.5 ± 1.01,500 ± 780

Fluorescent Toxin Reports Channel Activation.

We next tested whether we could optically measure channel activity with VST–fluorophore conjugates. The optical reporting ability of a GxTX–fluorophore was tested on CHO cells expressing Kv2.1 in whole-cell voltage-clamp fluorometry experiments. GxTX-dy550 was chosen for these experiments because it was found to have electrophysiological properties similar to GxTX (Fig. 1I and Fig. S2). To optically measure VST binding to voltage-clamped cells, we quantitated the fluorescence that remained after application and washout of 100 nM GxTX-dy550 (Fig. 4A). When cells were held at −100 mV, fluorescence slowly decayed. When cells were depolarized to 0 mV to accelerate toxin dissociation, fluorescence loss markedly accelerated (Fig. 4B). By fitting an exponential function (Eq. S4) to the fluorescence decays, rates of fluorescence change, k100 mV and k0 mV, were extracted (Fig. 4C). The similar kinetics of k100 mV fluorescence decay and koff of GxTX-dy550 at −100 mV (Fig. S2C) is consistent with fluorescence decay resulting from VST dissociating from channels. A clearly significant difference in fluorescence decay was seen after voltage change, with k0 mV being 53-fold faster than k100 mV. We conclude that the voltage-driven fluorescence change results from GxTX dissociation due to Kv2.1 voltage activation.
Fig. 4.
Fluorescent tarantula toxin reports Kv channel activity in Kv2.1-expressing CHO cells. (A) Fluorescence from a patch-clamped cell after washout of 100 nM GxTX-dy550. Bright-field (grayscale); GxTX-dy550 (red). (B) GxTX-dy550 fluorescence decay of voltage-clamped cell as in A. Holding potential switch from −100 mV to 0 mV indicated by blue bar. Gray line is fit of Eq. S4; k100 mV = 0.00399 ± 0.00001 s−1. Solid blue line is fit of Eq. S4 to initial 20 s at 0 mV; k0 mV = 0.166 ± 0.002 s−1. Dashed blue line is fit of Eq. S5; k0 mV fast = 0.222 ± 0.003 s−1, k0 mV slow = 0.0407 ± 0.0008 s−1. (C) Summary of dissociation rates of GxTX-dy550 from cells when held at −100 mV (black) or 0 mV (blue). Mean (solid line) ± SEM (dotted line). *P = 0.008. (D) Voltage-clamped (solid circle) and -unclamped (dashed circle) CHO cells bathed in 10 nM GxTX-dy550. Voltage of clamped cell held at −100 mV and stepped to 0 mV from 0 to 20 s. (E) Dotted black line, fluorescence change of unclamped cell: dashed circle in D. Solid black line, fluorescence change of voltage-clamped cell: solid circle in D. Blue bar indicates depolarization of clamped cell to 0 mV. Blue line is fit of Eq. S4, kF 0 mV = 0.163 ± 0.002 s−1. Gray line is fit of GxTX-dy550 reassociation at −100 mV, kF −100 mV = 0.0134 ± 0.0001 s−1. (F) Summary of kF in 10 nM GxTX-dy550. Line is fit of Eq. S6. V1/2 = -25 ± 11 mV, z = 1.5 ± 0.9e+, kF min = 0.011 ± 0.001 s, kF max = 0.15 ± 0.04 s. (G) Solid black line, fluorescence change of voltage-clamped cell in 1 nM GxTX-dy550. Holding potential = −80 mV. Blue bars indicate action-potential-like epochs: 2-ms steps to +40 mV at 100 Hz. Blue lines are fits of Eq. S4 during action-potential epochs; baseline (F0) constrained to the mean of final 10 s of last epoch (0.531 ∆F/F); kF = (in chronological order) 0.024 ± 0.005, 0.029 ± 0.001, and 0.071 ± 0.008 s−1.
To test whether Kv2.1 activation can be reversibly reported by this fluorescent VST, we bathed cells continuously in GxTX-dy550 and switched back and forth between voltages. In 10 nM GxTX, fluorescence was concentrated on the surface of Kv2.1-expressing cells (Fig. 4D) and remained constant over time (Fig. 4E) as the VST slowly bound to and dissociated from Kv2.1 channels. When voltage-clamped cells were held at a constant voltage, fluorescence intensity also remained constant. Varying voltage resulted in dynamic changes of cell fluorescence, indicating that the fluorescent VST optically reported the change in activation state of the ion channel (Fig. 4 D and E and Movie S1). The rate of fluorescence change (kF) varied with voltage and saturated at positive voltages (Fig. 4F), consistent with GxTX localization reporting activation of Kv channels rather than the membrane voltage. Thus, the activity of VGICs can be reported by state-selective fluorescent VST localization to live cells.
To test whether our VST probe can report channel activation at trace concentrations, which inhibit a negligible fraction of channels, we measured fluorescence from cells bathed in probe at a concentration substantially below the Kd of its target VGIC. At 1 nM, GxTX-dy550 is 30-fold below its measured Kd for Kv2.1, and has little effect on the whole-cell K+ current (Fig. S2A). At this inefficacious concentration, fluorescence could be detected on Kv2.1-expressing cells. This fluorescence was modulated by epochs of action-potential-like stimuli: 2-ms steps to +40 mV from a holding potential of −80 mV at 100 Hz. During these stimulus epochs, fluorescence decayed toward a similar baseline; in the periods between stimulus epochs, fluorescence slowly recovered (Fig. 4G). These signals suggest that every action potential increases the odds of probe dissociation. The mean fluorescence decay rate during the initial stimulus epoch was kF = 0.025 ± 0.005 s−1 (fit with Eq. S4, F0 unconstrained, n = 5 cells), intermediate to the dissociation rates seen at polarized or depolarized voltages (Fig. 4 C and F and Fig. S2C). This suggests that fluorescence change reports VGIC response to many action-potential-like stimuli integrated over time. The probe redistributes between channels and solution to reach a binding equilibrium dictated by the VGIC activation pattern. In this way, the state-dependent probe reports channel activation in response to action-potential-like voltage changes.

Channel-Modulating Toxins Mediate Ion Channel Binding to Microbeads.

The conformation-specific binding of GxTX–fluorophores to Kv channels suggests that the VST could guide other visible probes similarly. To determine if GxTX could imbue very large molecules with its binding properties, we tested whether cells expressing Kv2.1 channels would bind to 100-µm microbeads coated with GxTX. We chose beads used for one-bead–one-compound (OBOC) combinatorial drug discovery programs (4951) to establish whether GxTX-like molecules might be discoverable in an OBOC screen. These OBOC beads are composed of a solid-phase polystyrene dendrimer core with a PEG shell for water solubility and cell compatibility. We used copper(I)-catalyzed azide-alkyne Click cycloaddition to conjugate an alkyne-functionalized GxTX variant to azide-functionalized beads (Fig. 5A), as this chemistry successfully produced bioactive, PEGylated GxTX (Fig. 1E and Fig. S1 B and D). Beads with a GxTX coating adhered to Kv2.1-expressing cells (Fig. 5B). We found that bead–cell adhesion required both the VST peptide and a VGIC. Kv2.1 cells did not adhere well to control beads without GxTX, and cells lacking Kv2.1 did not adhere well to GxTX beads (Fig. 5 C and D). We next tested whether channel conformation impacted this GxTX-bead–Kv2.1-cell interaction. As studying adhesion of voltage-clamped cells to beads was infeasible, external [K+] was used to change voltage and activate channels. We first confirmed that increasing external [K+] depolarized cells to release GxTX from Kv2.1. Fluorescence decay resulting from GxTX-dy550 dissociation was measured in a control (5 mM) [K+] solution and after addition of a high (180 mM) [K+] solution. The high [K+] solution accelerated GxTX dissociation (Fig. 5 E and F), consistent with the high [K+] increasing resting voltage to activate the Kv2.1 channels. To assess the effects of channel activation on cell adhesion to GxTX-beads, Kv2.1-expressing cells were mixed with beads and allowed to adhere. These bead–cell complexes were then incubated in either a control or high [K+] solution. In blinded trials on five separate days, the fraction of beads with cells bound was reduced by incubation in high [K+] solution (Fig. 5G). Thus, the activity dependence of Kv2.1 binding was preserved when GxTX was attached to beads. The microbead results indicate that voltage sensor modulators can be identified by ion channel binding to OBOC beads. This suggests that new modulators of VGICs could be discovered in OBOC libraries. Taken together, these experiments show that VSTs can guide molecules of a wide range of sizes and compositions, from subnanometer fluorophores to 100-µm beads, and that these conjugates can act as reporters of VGIC activation.
Fig. 5.
Tarantula toxin-channel binding mediates cell adhesion to surfaces. (A) Depiction of GxTX Pra13 conjugation to azide beads and adhesion to cells. (B) Kv2.1 CHO cells adhering to a GxTX-coated bead. (C) Kv2.1 CHO cells selectively adhere to GxTX beads. Beads functionalized with either GxTX (gray) or fluorophore (blue) were incubated with CHO-K1 cells expressing Kv2.1. Note the propensity of cells to accumulate on GxTX beads. (D) Quantification of bead adhesion to cells as in A from experiments on separate days. Percentage of beads with more than four cells visibly adhered scored with GxTX on beads and Kv2.1 expressed on cells (gray, n = 6), with Kv2.1 cells but no GxTX on beads (green, n = 4), or with GxTX on beads but no Kv2.1 in cells (white, n = 3).*P < 0.05. (E) Fluorescence change of Kv2.1 CHO cells after washout of 100 nM GxTX. Blue bar indicates application of 7 volumes of high [K+] external. Lines are fits of Eq. S4; gray, kcontrol = 0.0069 ± 0.0002 s−1; blue, khigh[K+] = 0.0335 ± 0.0003 s−1. (F) Summary of dissociation rates of GxTX-dy550. *P = 0.008 (G) GxTX-coated beads lose adhesion to Kv2.1-expressing CHO cells in a depolarizing high [K+] solution. Each connected set of points represents time-matched samples from an experiment conducted on a different day. Experimenter was blinded to solutions used. **P < 0.05 Wilcoxon signed-rank directional test.


The functional roles of ion channels in biological circuits are actively debated. We optically detected activity of the Kv2.1 channel subtype using VST-conjugate probes. Kv2.1 is ubiquitously expressed in the soma of neurons throughout the brain (52) and is the dominant delayed rectifier of central neurons (53, 54). The channel’s activity shapes the repetitive firing properties of neurons (13), and Kv2.1 is homeostatically regulated in response to electrical activity (55). The up-regulation of Kv2.1 activity is proposed to relieve excitotoxic stress by suppressing neuronal hyperexcitability and seizure activity (32). However, under pathological conditions such as ischemia from stroke, up-regulation of Kv2.1 activity leads to apoptosis (56). To disentangle the contrasting neuroprotective and apoptotic roles of Kv2.1, it is essential to know when and to what degree the channel is activated. Activity-sensing probes such as the GxTX–fluorophores described here can detect channel response to sustained depolarization or epochs of action-potential-like stimuli (Fig. 4), and could provide a means to determine the extent to which Kv2 channels are involved with specific neural signaling pathways.
The full potential of ion channel activity probes would be realized with probes customized for particular channel types and specific questions related to their function. Developing activity probes for VGIC types requires careful consideration of their two components: a reporter amenable to sensitive detection and a conformation-specific VGIC ligand.
The ability of these probes to report channel activity without disrupting electrical signals is limited by the signal-to-noise characteristics of the reporter. An important caveat to consider when using any activity-dependent probe is the physics of microscopic reversibility, which dictates that a state-dependent ligand will act allosterically to modulate channel function. Similar limitations affect all other affinity-based molecular probes. For instance, calcium dyes buffer calcium and voltage-dependent dyes increase capacitance. To minimize perturbation of the process studied, probes are ideally used at a trace concentration. To avoid altering electrical signaling, VST probe can be applied at a concentration substantially below the Kd of its target VGIC as in Fig. 4G. At these electrophysiologically inefficacious concentrations, only a small fraction of channels is bound by the probe, yet optical signals from these probed channels can be detected. As the fraction of channel bound by a probe drops, the reporter signal diminishes, and hence a sensitive signal detection technique such as fluorescence microscopy is required. Reporters with increased signal-to-noise ratios would enable use of even lower trace concentrations of probe. Even brighter optical reporters such as quantum dots or up-converting nanoparticles (57) could enhance the VGIC activity-reporting capabilities of VST probes.
With a library of channel-specific optical probes, many secrets of ion channel activity could be rapidly revealed. This library could be engineered from the terrific variety of potent, VGIC subtype-selective VSTs that are increasingly being discovered. Examples of VSTs from spiders include ω-Agatoxin-IVA, which binds P-type Ca2+ channels (58) and the ProTx-I and -II peptides, which bind Na+ channel subtypes (5962), and heteropodatoxins, which bind Kv4 channels (63). The library of known VSTs is growing rapidly as venom peptides and mRNAs of more predatory animals are sequenced (64). Probes for different channel activities could be developed from toxins that stabilize different conformations of ion channels. For example, the Magi5 peptide stabilizes activated Na+ channels, whereas ProTx-II stabilizes resting conformations (65, 66). Similarly, GxTX inhibits K+ channels that are activated by the related VST, hanatoxin (35). All of these tarantula VSTs share a common fold, suggesting that each of them could be derivatized with chemoselective side chains, as was GxTX. Fluorescent labeling of different VSTs could create a palette of probes to report activation and deactivation of Ca2+, Na+, and K+ channel subtypes.
In addition, our results suggest a novel method of discovering activity-dependent ligands for VGICs. A clear difference was seen in binding between Kv2.1 cells and GxTX beads when the channels were voltage-activated (Fig. 5). This suggests that conformation-selective ligands could be found by screening combinatorial libraries using the OBOC method. In principle, an OBOC approach could be used to develop optical probes to report activity of different VGIC subtypes.
Specific VGIC activity-sensing probes also have the potential to map ion channel activation throughout large regions of the brain and body. Using these probes in intact tissue could enable measurement of which channel subtypes respond to different stimuli within biological circuits. Our laboratories are currently developing probes with improved optical and kinetic properties to report activation of ion channels in vivo.


This study describes optical reporting of ion channel activation in live cells. Our study demonstrates that: (i) voltage sensor toxins can retain bioactivity after chemoselective conjugation to reporters, (ii) fluorescent toxins colocalize with channels in live cells, (iii) toxin fluorescence reports channel activation, and (iv) toxins mediate activity-dependent binding of channels to microbeads. The activation-state-dependent binding of these reporters to live cells suggests potential for dissecting specific contributions of ion channels to cellular electrical activity and enabling discovery of novel ion channel modulators.

Materials and Methods

Details are in SI Materials and Methods. Peptides were synthesized by Fmoc peptide synthesis, folded by air oxidation, and purified by reverse-phase HPLC. Identity of peptides and conjugates was verified by matrix-assisted laser desorption/ionization time of flight (MALDI-TOF) mass spectrometry, HPLC, and SDS/PAGE. Whole-cell voltage-clamp recording was used to measure currents from CHO-K1 cells stably expressing rat Kv2.1 channels. Live cell imaging and patch-clamp fluorometry were performed on an inverted microscope using light-emitting diode or laser excitation. Structural modeling was performed using the Rosetta-Membrane method (3638).


We thank Yi Liu of the Molecular Foundry for the BTTAA reagent. We are very grateful to many at the University of California, Davis: Oscar Cerda for his talented culturing of hippocampal neurons; James Trimmer for ion channel plasmids and helpful discussion; Jie Zheng and Peter Cala for feedback on the manuscript; and Sebastian Ayala, Christina Berry, and Yuanpei Li for providing technical assistance. This work was supported by US NIH Grants 5P30GM092328-02, R01NS042225-09S1, R25NS063307, and T32HL086350; American Heart Association Grant 10SDG4220047; and Milton L. Shifman Endowed Scholarship for the Neurobiology Course at Woods Hole. Work at the Molecular Foundry was supported by the Office of Science, Office of Basic Energy Sciences, of the US Department of Energy under Contract DE-AC02-05CH11231. This work is dedicated to the memory of coauthor Kenneth S. Eum (1987–2014). Ken was a talented PhD student, a driven and caring soul who brought joy to the lives of those who knew him.

Supporting Information

Supporting Information (PDF)
Supporting Information


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Information & Authors


Published in

Go to Proceedings of the National Academy of Sciences
Proceedings of the National Academy of Sciences
Vol. 111 | No. 44
November 4, 2014
PubMed: 25331865


Submission history

Published online: October 20, 2014
Published in issue: November 4, 2014


  1. voltage-gated ion channel
  2. potassium channel
  3. gating modifier
  4. fluorescence
  5. allostery


We thank Yi Liu of the Molecular Foundry for the BTTAA reagent. We are very grateful to many at the University of California, Davis: Oscar Cerda for his talented culturing of hippocampal neurons; James Trimmer for ion channel plasmids and helpful discussion; Jie Zheng and Peter Cala for feedback on the manuscript; and Sebastian Ayala, Christina Berry, and Yuanpei Li for providing technical assistance. This work was supported by US NIH Grants 5P30GM092328-02, R01NS042225-09S1, R25NS063307, and T32HL086350; American Heart Association Grant 10SDG4220047; and Milton L. Shifman Endowed Scholarship for the Neurobiology Course at Woods Hole. Work at the Molecular Foundry was supported by the Office of Science, Office of Basic Energy Sciences, of the US Department of Energy under Contract DE-AC02-05CH11231. This work is dedicated to the memory of coauthor Kenneth S. Eum (1987–2014). Ken was a talented PhD student, a driven and caring soul who brought joy to the lives of those who knew him.


*This Direct Submission article had a prearranged editor.



Drew C. Tilley1
Department of Physiology and Membrane Biology, University of California, Davis, CA 95616;
Kenneth S. Eum1,2
Department of Physiology and Membrane Biology, University of California, Davis, CA 95616;
Neurobiology Course, Marine Biological Laboratory, Woods Hole, MA 02543;
Sebastian Fletcher-Taylor
Department of Physiology and Membrane Biology, University of California, Davis, CA 95616;
Daniel C. Austin
Department of Physiology and Membrane Biology, University of California, Davis, CA 95616;
Christophe Dupré
Neurobiology Course, Marine Biological Laboratory, Woods Hole, MA 02543;
Lilian A. Patrón
Neurobiology Course, Marine Biological Laboratory, Woods Hole, MA 02543;
Rita L. Garcia
Molecular Foundry, Lawrence Berkeley National Laboratory, Berkeley, CA 94720; and
Kit Lam
Department of Biochemistry and Molecular Medicine, University of California, Davis, CA 95616
Vladimir Yarov-Yarovoy
Department of Physiology and Membrane Biology, University of California, Davis, CA 95616;
Department of Biochemistry and Molecular Medicine, University of California, Davis, CA 95616
Bruce E. Cohen
Molecular Foundry, Lawrence Berkeley National Laboratory, Berkeley, CA 94720; and
Department of Physiology and Membrane Biology, University of California, Davis, CA 95616;
Neurobiology Course, Marine Biological Laboratory, Woods Hole, MA 02543;


To whom correspondence should be addressed. Email: [email protected].
Author contributions: D.C.T., K.S.E., V.Y.-Y., B.E.C., and J.T.S. designed research; D.C.T., K.S.E., D.C.A., C.D., L.A.P., and J.T.S. performed research; S.F.-T., R.L.G., K.L., B.E.C., and J.T.S. contributed new reagents/analytic tools; D.C.T., K.S.E., D.C.A., C.D., L.A.P., and J.T.S. analyzed data; and D.C.T., K.S.E., V.Y.-Y., B.E.C., and J.T.S. wrote the paper.
D.C.T. and K.S.E. contributed equally to this work.
Deceased June 22, 2014.

Competing Interests

The authors declare no conflict of interest.

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    Chemoselective tarantula toxins report voltage activation of wild-type ion channels in live cells
    Proceedings of the National Academy of Sciences
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