Major diversification of voltage-gated K⁺ channels occurred in ancestral parahoxozoans

Significance

We examined the origin and evolution of two major families of voltage-gated K⁺ channels, Shaker and KCNQ, which regulate action potential repolarization, patterning, and threshold. Shaker family channels evolved in a basal metazoan ancestor of ctenophores and parahoxozoans (including cnidarians and bilaterians), but functional diversification of the Shaker family and the emergence of the KCNQ family occurred specifically within the parahoxozoan lineage. Our results suggest that many major innovations in the regulation of cellular excitability by voltage-gated K⁺ channels are unique to parahoxozoans and that these innovations occurred before the divergence of cnidarians and bilaterians. Ctenophores and sponges separated prior to this burst of innovation and thus either lack major mechanisms for action potential regulation or evolved such mechanisms independently.

Abstract

We examined the origins and functional evolution of the Shaker and KCNQ families of voltage-gated K⁺ channels to better understand how neuronal excitability evolved. In bilaterians, the Shaker family consists of four functionally distinct gene families (Shaker, Shab, Shal, and Shaw) that share a subunit structure consisting of a voltage-gated K⁺ channel motif coupled to a cytoplasmic domain that mediates subfamily-exclusive assembly (T1). We traced the origin of this unique Shaker subunit structure to a common ancestor of ctenophores and parahoxozoans (cnidarians, bilaterians, and placozoans). Thus, the Shaker family is metazoan specific but is likely to have evolved in a basal metazoan. Phylogenetic analysis suggested that the Shaker subfamily could predate the divergence of ctenophores and parahoxozoans, but that the Shab, Shal, and Shaw subfamilies are parahoxozoan specific. In support of this, putative ctenophore Shaker subfamily channel subunits coassembled with cnidarian and mouse Shaker subunits, but not with cnidarian Shab, Shal, or Shaw subunits. The KCNQ family, which has a distinct subunit structure, also appears solely within the parahoxozoan lineage. Functional analysis indicated that the characteristic properties of Shaker, Shab, Shal, Shaw, and KCNQ currents evolved before the divergence of cnidarians and bilaterians. These results show that a major diversification of voltage-gated K⁺ channels occurred in ancestral parahoxozoans and imply that many fundamental mechanisms for the regulation of action potential propagation evolved at this time. Our results further suggest that there are likely to be substantial differences in the regulation of neuronal excitability between ctenophores and parahoxozoans.

Voltage-gated K⁺ channels are highly conserved among bilaterian metazoans and play a central role in the regulation of excitation in neurons and muscle. Understanding the functional evolution of these channels may therefore provide important insights into how neuromuscular excitation evolved within the Metazoa. Three major gene families, Shaker, KCNQ, and Ether-a-go-go (EAG) encode all voltage-gated K⁺ channels in bilaterians (1, 2). In this study, we examine the functional evolution and origins of the Shaker and KCNQ gene families. Shaker family channels can be definitively identified by a unique subunit structure that includes both a voltage-gated K⁺ channel core and a family-specific cytoplasmic domain within the N terminus known as the T1 domain. T1 mediates assembly of Shaker family subunits into functional tetrameric channels (3, 4). KCNQ channels are also tetrameric but lack a T1 domain and use a distinct coiled-coil assembly domain in the C terminus (5, 6). KCNQ channels can be identified by the presence of this family-specific assembly motif and high amino acid conservation within the K⁺ channel core. Both channel families are found in cnidarians (1, 7) and thus predate the divergence of cnidarians and bilaterians, but their ultimate evolutionary origins have not yet been defined.

Shaker family K⁺ channels serve diverse roles in the regulation of neuronal firing and can be divided into four gene subfamilies based on function and sequence homology: Shaker, Shab, Shal, and Shaw (8, 9). The T1 assembly domain is only compatible between subunits from the same gene subfamily (4, 10) and thus serves to keep the subfamilies functionally segregated. Shaker subfamily channels activate rapidly near action potential threshold and range from rapidly inactivating to noninactivating. Multiple roles for Shaker channels in neurons and muscles have been described, but their most unique and fundamental role may be that of axonal action potential repolarization. Shaker channels are clustered to the axon initial segment and nodes of Ranvier in vertebrate neurons (11–13) and underlie the delayed rectifier in squid giant axons (14). The Shaker subfamily is diverse in cnidarians (15, 16), and the starlet sea anemone Nematostella vectensis has functional orthologs of most identified Shaker current types observed in bilaterians (16).

The Shab and Shal gene subfamilies encode somatodendritic delayed rectifiers and A currents, respectively (17–20). Shab channels are important for maintaining sustained firing (21, 22), whereas the Kv4-based A current modulates spike threshold and frequency (17). Shab and Shal channels are present in cnidarians, but cnidarian Shab channels have not been functionally characterized, and the only cnidarian Shal channels expressed to date display atypical voltage dependence and kinetics compared with bilaterian channels (23). Shaw channels are rapid, high-threshold channels specialized for sustaining fast firing in vertebrates (24, 25) but have a low activation threshold and may contribute to resting potential in Drosophila (19, 26, 27). A Caenorhabditis elegans Shaw has slow kinetics but a high activation threshold (28), and a single expressed cnidarian Shaw channel has the opposite: a low activation threshold but relatively fast kinetics (29). Thus, the ancestral properties and function of Shaw channels is not yet understood. Further functional characterization of cnidarian Shab, Shal, and Shaw channels would provide a better understanding of the evolutionary status of the Shaker family in early parahoxozoans.

KCNQ family channels underlie the M current in vertebrate neurons (30) that regulates subthreshold excitability (31). The M current provides a fundamental mechanism for regulation of firing threshold through the Gq G-protein pathway because KCNQ channels require phosphatidylinositol 4,5-bisphosphate (PIP₂) for activation (32, 33). PIP₂ hydrolysis and subsequent KCNQ channel closure initiated by Gq-coupled receptors produces slow excitatory postsynaptic potentials, during which the probability of firing is greatly increased (32, 33). The key functional adaptations of KCNQ channels for this physiological role that can be observed in vitro are (i) a requirement for PIP₂ to couple voltage-sensor activation to pore opening (34, 35), and (ii) a hyperpolarized voltage–activation curve that allows channels to open below typical action potential thresholds. Both key features are found in vertebrate (30, 34, 36–38), Drosophila (39), and C. elegans (40) KCNQ channels, suggesting they may have been present in KCNQ channels in a bilaterian ancestor. Evolution of the M current likely represented a major advance in the ability to modulate the activity of neuronal circuits, but it is not yet clear when PIP₂-dependent KCNQ channels first evolved.

Here, we examine the origins and functional evolution of the Shaker and KCNQ gene families. If we assume the evolution of neuronal signaling provided a major selective pressure for the functional diversification of voltage-gated K⁺ channels, then we can hypothesize that the appearance of these gene families might accompany the emergence of the first nervous systems or a major event in nervous system evolution. Recent phylogenies that place the divergence of ctenophores near the root of the metazoan tree suggest that the first nervous systems, or at least the capacity to make neurons, may have been present in a basal metazoan ancestor (41–43) (Fig. S1). One hypothesis then is that much of the diversity of metazoan voltage-gated channels should be shared between ctenophores and parahoxozoans [cnidarians, bilaterians, and placozoans (44)]. However, genome analysis indicates that many “typical” neuronal genes are missing in ctenophores and the sponges lack a nervous system, leading to the suggestion that extant nervous systems may have evolved independently in ctenophores and parahoxozoans (42, 45). Thus, a second hypothesis is that important steps in voltage-gated K⁺ channel evolution might have occurred separately in ctenophores and parahoxozoans. We tested these hypotheses by carefully examining the phylogenetic distribution and functional evolution of Shaker and KCNQ family K⁺ channels. Our results support a model in which major innovations in neuromuscular excitability occurred specifically within the parahoxozoan lineage.

Results

To determine the evolutionary origin of the Shaker family, we used a BLAST (46) search strategy to identify potential Shaker family genes in choanoflagellates, ctenophores, and sponge with bilaterian and cnidarian Shaker, Shab, Shal, and Shaw proteins as queries. We defined Shaker family channels as (i) including a T1 domain, and (ii) having best matches to known Shaker family channels within the voltage-gated K⁺ channel core motif in reciprocal BLAST searches against bilaterian and cnidarian sequences. Shaker family channels were not present in genome drafts and gene predictions from two choanoflagellates, Salpingoeca rosetta and Monosiga brevicollis (47, 48). Because choanoflagellates are believed to represent the closest extant relatives of metazoans, this is a strong indication that the Shaker family is metazoan specific. In support of this view, voltage-gated K⁺ channels cloned from prokaryotes (49), plants (50, 51), and fungi (52) lack the T1 domain and Shaker-specific homology.

We next searched for Shaker family channels in sponges and ctenophores because current views of metazoan phylogeny place ctenophores or sponges as the most basally branching extant phyla (41–43). We were unable to find Shaker channels in the genome of the sponge Amphimedon queenslandica (53), or in seven of eight sponge species transcriptomes (54). However, three ESTs from the Corticium candelabrum transcriptome could be assembled into two nonoverlapping fragments with specifically high homology to the Shaker subfamily, one covering T1–S1 and one covering S3 to the C terminus (Fig. S2). In contrast, we identified 49 Shaker family genes in the draft genome and transcriptome of the ctenophore Mnemiopsis leidyi (41). Amino acid predictions for channel proteins encoded by these genes are included in Dataset S1. Multiple Shaker family channels were also present in the genome draft of a second ctenophore, Pleurobachia bachei, and four ctenophore transcriptomes (Pleurobachia plus three additional species) (42). However, we did not assemble Shaker channel sets from these species because many Pleurobachia gene predictions and transcriptome sequences were fragmentary. As a whole, these results indicate that the Shaker family originated in basal metazoans. The reason for the apparent absence of Shaker family channels in many sponges is currently unclear but could point to loss of the family within various sponge lineages.

To determine the evolutionary relationship between ctenophore and parahoxozoan Shaker family channels, we constructed a Bayesian inference phylogeny (Fig. 1) based on alignment of the T1 and voltage-gated K⁺ channel core motifs. We excluded seven ctenophore channels and the Corticium candelabrum fragments because of large sequence gaps in these motifs. Each of the seven excluded ctenophore channels had highest homology to other Mnemiopsis sequences in BLAST comparisons. The phylogeny splits Mnemiopsis Shaker family channels into two clades, one separate from the parahoxozoan channels and one within the parahoxozoan Shaker subfamily (Fig. 1). The phylogeny therefore supports a model in which the Shaker subfamily evolved before the divergence of ctenophores and parahoxozoans, and that Shab, Shal, and Shaw later evolved from Shaker specifically within the parahoxozoan lineage. All top hits identified in BLAST searches of Pleurobrachia bachei (42) using Shab, Shal, and Shaw channels best matched identified Mnemiopsis Shaker family sequences when used as queries in reciprocal BLAST searches. Therefore, both ctenophores species appear to lack Shab, Shal, and Shaw channels. We rebuilt and tested the phylogeny using both minimum-evolution and maximum-likelihood methods to further assess these findings. Minimum evolution supports the same topology as Bayesian inference (Fig. S3). Maximum likelihood supports the late evolution of Shab, Shal, and Shaw and a close relationship between one ctenophore channel clade and the Shaker subfamily, but is unable to resolve the branching pattern (Fig. S3) between them. Sequence similarity network (SSN) analysis also agrees with the Bayesian inference topology (Fig. S4). These sequence analyses therefore converge on a model in which the Shaker subfamily evolved before the Shab, Shal, and Shaw subfamilies and possibly before the ctenophore/parahoxozoan divergence.

Fig. 1.

Bayesian inference phylogeny of the metazoan Shaker K⁺ channel family. Major metazoan clades are identified by color according to the legend at the Upper Left; the Shaker, Shab, Shal, and Shaw subfamilies are marked at the right margin, and protein names are given at branch termini. Species prefixes in the protein names in alphabetical order are as follows: C.ele (Caenorhabditis elegans, nematode), D.mel (Drosophila melanogaster, fruit fly), H.sap (Homo sapiens, human), M.lei (Mnemiopsis leidyi, ctenophore), N.vec (Nematostella vectensis, sea anemone), S.pur (Stronglyocentrotus purpuratus, sea urchin), and T.adh (Trichoplax adhaerens, placozoan). Green asterisks mark channels functionally expressed in this study. Sequences used in phylogeny construction are provided in Dataset S1. The scale bar indicates number of substitutions per site, and posterior probabilities are given at branch nodes. Unlabeled nodes had posterior probabilities ≥0.97.

Nematostella Shaker channels have been extensively characterized and demonstrate functional conservation between hydrozoans, anthozoans, and bilaterians (15, 16, 55). To better understand the functional evolution of the Shaker family, we first functionally expressed Nematostella whole Shab, Shaw, and Shal channels in Xenopus oocytes. Cnidarian Shab channels have not previously been expressed. Nematostella Shab (NvShab) currents are compared with the mouse Shab channel Kv2.1 in Fig. 2A. They encode highly similar delayed rectifier channels, although conductance–voltage and steady-state inactivation relationships (Fig. 2 B and C) reveal a modest hyperpolarized shift in the voltage dependence of NvShab. V₅₀ and slope values for the single-Boltzmann fits of voltage–activation and steady-state inactivation curves are included in Table 1. These classic delayed rectifier properties coupled with partial steady-state inactivation are shared among vertebrate, Drosophila (26), and Nematostella Shab currents. In contrast, an Aplysia Shab channel inactivates within a few hundred milliseconds (56) and a nematode Shab family channel EXP-2 encodes a rapidly inactivating, I_Kr-like current (57). Both lophotrochozoans and nematodes have large Shab subfamily gene expansions (16) that remain functionally uncharacterized, so the full diversity of Shab currents in these species is not currently known. Our results suggest that the delayed rectifier Shab phenotype was present before the cnidarian/bilaterian divergence and probably represents the ancestral functional phenotype of the gene subfamily.

Fig. 2.

NvShab encodes a typical Shab family delayed rectifier. (A) Examples of NvShab and mouse Kv2.1 current traces elicited by 400-ms voltage steps from −50 to 50 mV in 20-mV increments (holding at −100 mV, tail at −50 mV). (B) Normalized conductance–voltage (GV) relationships measured from isochronal tail currents following 400-ms voltage steps to the indicated voltages. (C) Steady-state inactivation measured from peak current during test pulse to 10 mV following 4.5-s prepulses to the indicated voltages (holding at −100 mV). Curves in B and C show single-Boltzmann distribution fits (parameters in Table 1); error bars show SEM, n = 7–8.

Table 1.

Boltzmann fit parameters for Shaker and KCNQ channel family GV and steady-state inactivation (SSI) curves

Channel	n	V50, mV	s, mV
GV
NvShab	7	−13.4 ± 1.4	12.8 ± 0.6
MmKv2.1	8	−1.0 ± 1.6	14.3 ± 0.4
NvShaw1	6	−10.6 ± 5.0	22.1 ± 1.0
NvShaw1 + R1	8	75.0 ± 1.8	27.2 ± 1.1
NvShaw1 + R2 (first, fraction 0.09 ± 0.02)	9	10.9 ± 0.5	6.6 ± 0.6
NvShaw1 + R2 (second)	9	82.1 ± 1.8	17.6 ± 0.2
NvShal1	11	18.2 ± 1.2	15.3 ± 0.6
NvShal1 + R1	8	−11.2 ± 1.7	14.6 ± 0.4
NvShal1 + R2	12	−12.7 ± 1.5	16.2 ± 0.3
NvShal1 + R3	7	−3.0 ± 2.2	15.2 ± 0.6
NvKCNQ	10	−36.6 ± 1.6	8.6 ± 0.2
HsKCNQ2/3	14	−13.6 ± 1.5	20.6 ± 0.6
SSI
NvShab	7	−34.9 ± 0.5	5.1 ± 0.2
MmKv2.1	7	−8.8 ± 0.7	8.6 ± 0.5
NvShal1	9	−33.1 ± 0.8	6.9 ± 0.3
NvShal1 + R1	6	−20.0 ± 0.8	11.6 ± 0.7
NvShal1 + R2	8	−71.7 ± 1.2	7.8 ± 0.1
NvShal1 + R3	9	−63.6 ± 1.3	13.1 ± 0.3

n, number of samples; V₅₀, half-maximal activation voltage; s, slope factor.

We next characterized the functional properties of several NvShaw channels. The Nematostella Shaw subfamily comprises 11 genes (Fig. 1). We first expressed one of two highly conserved (with respect to bilaterians) channels at the base of the Nematostella expansion, NvShaw1. Like NvShab, NvShaw1 expressed functional homomeric channels in Xenopus oocytes; currents had a lower activation threshold and shallower voltage dependence compared with NvShab (Fig. 3 A and C, and Table 1). NvShaw1 currents are similar to Drosophila Shaw (26) and a Shaw channel from the hydrozoan Polyorchis penicillatus (29). We failed to obtain functional homomeric channels when we expressed two additional Shaw channels from the heart of the Nematostella gene expansion (Fig. 3A); we postulated that these two channels, which we named NvShawR1 and NvShawR2, may encode regulatory subunits that only assemble into functional heteromeric channels. Eleven of 17 expressed Nematostella Shaker channels have this regulatory phenotype (16). Coexpression of NvShawR1 and NvShawR2 with NvShaw1 indeed produced heteromeric currents with slower activation kinetics and a strongly depolarized activation threshold compared with NvShaw1 alone (Fig. 3 B and C, and Table 1). The NvShaw1+NvShawR2 voltage–activation curve was best fit by a double Boltzmann, suggesting the presence of two functionally distinct heteromers that presumably differ by stoichiometry. These heteromeric currents are similar to the C. elegans Shaw gene egl-36 (28) but differ from high-threshold vertebrate Shaw currents by their slower kinetics and shallow voltage dependence. The regulatory subunit phenotype has been found in vertebrate Shab channels (58, 59), hydrozoan Shal channels (23), and Nematostella Shaker channels (16), but has not previously been described for the Shaw subfamily.

Fig. 3.

Functional properties of homomeric and heteromeric Nematostella Shaw channels. (A) Current traces elicited from oocytes injected with NvShaw1, NvShawR1, and NvShawR2 by 400-ms voltage steps from −60 to 60 mV in 20-mV increments (holding at −100 mV; tail at −50 mV). (B) Superimposed current traces from oocytes injected with NvShaw1 alone (gray), or NvShaw1+NvShawR1 (Left) and NvShaw1+NvShawR2 (Right). Traces were normalized to peak current at 60 mV for comparison. (C) Normalized GV curves of homomeric and heteromeric Nematostella Shaw channels. Data were taken from isochronal tail currents following 400-ms voltage steps for NvShaw1, and from peak current adjusted for driving force (assuming a −100-mV reversal potential) for heteromeric currents. Curves show single (NvShaw1 and NvShaw1+R1)- or double (NvShaw1+R2)-Boltzmann fits (parameters reported in Table 1; error bars show SEM, n = 6–9). NvShaw1 was injected at a 1:10 RNA ratio relative to NvShawR1 and NvShawR2 to produce a predominantly heteromeric current for comparison.

The Nematostella Shal subfamily has a single highly conserved gene (relative to bilaterians), NvShal1, which is an ortholog of the hydrozoan (Polyorchis) channel jShal1, and a Nematostella-specific expansion of 11 genes (Fig. 1 and ref. 16). The Polyorchis Shal regulatory subunit jShalγ1 is closely related to the Nematostella Shal expansion (16), suggesting that these genes may also encode subunits with a regulatory phenotype. NvShal1 expressed in Xenopus oocytes produce the classical transient A-type current, but with a high activation threshold (Fig. 4 A and B). Bilaterian Shal channels typically begin to activate around −40 to −50 mV, and jShal1 activation can be observed as low as −80 mV (23). However, C. elegans Shal has a similarly depolarized activation threshold (60). Although Shal currents vary in activation threshold, all Shal channels described to date have the classic A-current property of closed-state inactivation, which results in largely nonoverlapping steady-state inactivation and voltage–activation curves (61). Thus, the Shal-based A current is most active in neurons when the membrane has been hyperpolarized before a depolarization (17). NvShal1 shares this characteristic separation of voltage–activation and steady-state inactivation (Fig. 4B). In contrast, the partial steady-state inactivation of delayed rectifier Shab channels overlaps the voltage–activation curve (Fig. 2C). We cloned and expressed three Shal genes from the Nematostella expansion, NvShalR1–R3, and confirmed that each has the regulatory phenotype. NvShalR1–R3 did not produce outward K⁺ currents when expressed alone (Fig. 4C) but formed heteromeric channels with distinct properties when coexpressed with NvShal1 (Fig. 4 D–F). NvShalR1 hyperpolarized the GV curve and converted the phenotype of NvShal1 to that of a Shab-like delayed rectifier: inactivation was greatly reduced during short steps (Fig. 4D) and partial steady-state inactivation shifted to the Shab pattern of overlap with the voltage–activation curve (Fig. 4E and Table 1). This is the first example to our knowledge of a Shal channel that lacks an A-current phenotype. In contrast, NvShalR2 and NvShalR3 altered the inactivation rate observed in current traces (Fig. 4D) and hyperpolarized both the GV and steady-state inactivation curves (Fig. 4E and Table 1), but preserved the classic A-current phenotype. NvShalR2 and NvShalR3 also slow the rate of recovery from inactivation (Fig. 4F). These results indicate that the large Nematostella Shal subfamily encodes a variety of A currents and that at least one heteromeric Shal current (NvShal1+NvShalR1) instead contributes to delayed rectifier diversity, typically the domain of the Shab subfamily.

Fig. 4.

Functional properties of homomeric and heteromeric Nematostella Shal channels. (A) Current traces from an oocyte expressing NvShal1 elicited by 1-s voltage steps (−60 to 60 mV in 20-mV increments; holding at −100 mV and tail at −50 mV). (B) Normalized GV (black) and steady-state inactivation (gray) relationships for NvShal1. GV was measured from peak current during depolarizing steps and adjusted for driving force (assuming −100-mV reversal). Steady-state inactivation was measured from peak current at +40 mV following 4.5-s steps to the indicated voltages. (C) No K⁺ currents could be elicited from oocytes injected with NvShalR1-3; voltage protocols are the same as in A. (D) Current traces elicited by the same voltage protocols from oocytes injected with NvShal1+NvShalR1–R3 (1:10 RNA ratio). (E) Normalized GV (black) and steady-state inactivation (gray) relationships for heteromeric Nematostella Shal currents, measured and fit as in B. The dashed lines are Boltzmann fits of data from homomeric NvShal1 currents shown in B. Data in B and E show average values, and error bars indicate SEM (n = 6–12); curves in B and E show single-Boltzmann fits (V₅₀ and slope values reported in Table 1). (F) Time course of recovery from inactivation for NvShal1, NvShal1+NvShalR2, and NvShal1+NvShalR3. Recovery was determined using a double 1-s, +40-mV pulse protocol: data show fraction of current recovered in a second depolarizing pulse after a repolarizing step to −100 mV of the indicated duration. Curves show a double-exponential fit of the data. Time constants (τ) for NvShal1 were 7.8 ± 1.2 (0.68 ± 0.03 fractional amplitude) and 36.5 ± 6.3 (n = 10; ±SEM). Time constants for NvShal1+NvShalR2 were 74.4 ± 4.5 (0.60 ± 0.02 fractional amplitude) and 350.5 ± 30.5 (n = 14), and time constants for NvShal1+NvShalR3 were 98.0 ± 9.4 (0.25 ± 0.01 fractional amplitude) and 1,030.9 ± 119.2 (n = 8).

We next cloned two of the potential Mnemiopsis Shaker subfamily channels, MlShak1 and MlShak2, to examine their functional phenotypes in vitro. However, neither of the two channels produced functional voltage-gated potassium currents when expressed alone or in combination in Xenopus oocytes (Fig. 5A). We reasoned that the channels we chose to express could be regulatory subunits, but identifying viable heteromeric combinations of subunits among 49 Mnemiopsis Shaker family genes with no a priori knowledge of mixing rules was a daunting proposition. We therefore focused on functionally testing their putative Shaker subfamily identity suggested by phylogenetic analysis. If MlShak1 and MlShak2 are Shaker subfamily genes, then they should specifically coassemble with parahoxozoan Shaker subfamily channels. We therefore coexpressed them with Nematostella (NvShak3) and mouse (MmKv1.2) Shaker channels. RNAs were mixed at a 10:1 ratio in favor of Mnemiopsis subunits to bias formation of heteromers. Both ctenophore channels introduced a slowly activating component into NvShak3 currents and slowed channel deactivation (Fig. 5B), indicating that functional heteromers were formed. Currents were also significantly reduced by MlShak1 coexpression (Fig. 5D), suggesting either a smaller single-channel conductance for heteromers or that not all assembled channel stoichiometries were functional. When the ctenophore channels were coexpressed with mouse Kv1.2, this dominant-negative suppression was the major phenotype; currents were reduced by ∼90% (Fig. 5 C and E). We did not closely examine the small residual currents for evidence of functional heteromer formation; dominant-negative suppression is commonly used as evidence for coassembly of K⁺ channel subunits (62–64).

Fig. 5.

Ctenophore Shaker family subunits coassemble with Nematostella and mouse Shaker subfamily subunits. (A) No outward K⁺ currents were detected in oocytes expressing the Mnemiopsis leidyi Shaker family channels MlShak1 and MlShak2 either alone or in combination (1:1 RNA ratio) (1-s voltage steps from −60 to 60 mV in 20-mV increments, holding at −100 mV). (B and C) Current traces showing the Nematostella Shaker subfamily channel NvShak3 (B) and the mouse Shaker subfamily channel Kv1.2 (MmKv1.2; C) expressed alone or together with either MlShak1 or MlShak2 (1:10 RNA ratio). Currents were elicited by 400-ms voltage steps from −60 to 60 mV in 20-mV increments (holding at −100 mV; tail at 0 mV for NvShak3 and −50 mV for MmKv1.2). Gray traces in B show NvShak3 current at 60 mV normalized to the heteromeric channel peak current at 60 mV to illustrate changes in current shape; note the pronounced slowing of deactivation (arrows). (D and E) Comparison of average peak outward current amplitudes recorded at 60 mV for NvShak3 (D) and MmKv1.2 (E) alone and in combination with MlShak1 and MlShak2. Error bars show SEM, and n is indicated on the graphs. Asterisks indicate significant difference in amplitude from NvShak3 or MmKv1.2: *P < 0.05; ***P < 0.001 (two-tailed t test).

We further tested whether coassembly with MlShak1 and MlShak2 was specific to the Shaker subfamily by coexpressing the channels with NvShab, NvShal1, and NvShaw1. We did not observe novel components in the currents when these channels were mixed with MlShak1 and MlShak2 at RNA concentrations that specifically altered NvShak3 and MmKv1.2 currents (Fig. 6 A, C, and E). Furthermore, we did not observe dominant-negative suppression of currents during these coexpression experiments (Fig. 6 B, D, and F). These results therefore indicate that functional and dominant-negative assembly with MlShak1 and MlShak2 is limited to Shaker subfamily and thus support the hypothesis that MlShak1 and MlShak2 are Shaker subfamily channels.

Fig. 6.

Ctenophore Shaker family subunits do not coassemble with Nematostella Shab, Shal, or Shaw channels. (A) Current traces are compared for NvShab (gray) and NvShab+MlShak1 (Left, black) and NvShab+MlShak2 (Right, black). Currents were elicited by 1-s steps ranging from −60 to 60 mV in 20-mV increments and normalized to the amplitude at 60 mV. No differences in current shape were observed. (B) Peak outward current amplitudes at 60 mV are shown for NvShab, NvShab+MlShak1, and NvShab+MlShak2 (average of the indicated number of measurements; error bars show SEM). No significant differences in current amplitude were observed in two-tailed t test comparisons (n.s.). (C–F) Identical analyses are shown for NvShaw1 and NvShal1. NvShab, NvShaw1, and NvShal1 were coexpressed with MlShak1 and MlShak2 at a 1:10 RNA ratio.

We next examined the origins and functional evolution of the KCNQ gene family, starting with a similar BLAST search strategy. We defined KCNQ genes as containing the C-terminal coiled-coil assembly domains and having greater homology to KCNQ channels in voltage-gated K⁺ channel core in reciprocal BLAST searches against bilaterian sequences. We did not find KCNQ family genes outside the parahoxozoan lineage: There were no KCNQ family genes in two choanoflagellate genomes (47, 48), two ctenophore genomes and five ctenophore transcriptomes (41, 42), or one sponge genome and eight sponge transcriptomes (53, 54). Furthermore, we were unable to find KCNQ channels in the placozoan Trichoplax adhaerens (65), a parahoxozoan. However, KCNQ channels can be found in all cnidarian and bilaterian genomes we have examined. Anthozoans Nematostella vectensis, Acropora digitifera, and Orbicella faveolata had a single KCNQ channel, whereas the hydrozoan Hydra magnipapillata (66) had six KCNQ channels. Phylogenetic analysis indicated that two distinct clades of KCNQ channels are present in bilaterians and that all cnidarian KCNQ channels fall outside these clades (Fig. 7). The phylogeny supports a model in which a cnidarian/bilaterian ancestor had a single KCNQ channel, which was then duplicated in an ancestral bilaterian after divergence from cnidarians. Members of both bilaterian KCNQ clades display the characteristic PIP₂ dependence of the channel family (34), suggesting that this defining feature evolved before the duplication of KCNQ in bilaterians.

Fig. 7.

Bayesian inference phylogeny of the metazoan KCNQ voltage-gated K⁺ channel family. Color indicates major phylogenetic groups of metazoans, and dashed gray lines separate bilaterian KCNQ1-like, bilaterian KCNQ2-like, and cnidarian KCNQ sequences. Channel names include the following species prefixes: A.dig (Acropora digitifera, coral), A.gam (Anopheles gambiae, mosquito), C.ele (Caenorhabditis elegans, nematode), C.tel (Capitella teleta, polychaete), C.int (Ciona intestinalis, tunicate), C.gig (Crassostrea gigas, oyster), D.mel (Drosophila melanogaster, fruit fly), H.mag (Hydra magnipapillata, hydra), L.gig (Lottia gigantea, limpet), M.mus (Mus musculus, mouse), N.vec (Nematostella vectensis, sea anemone), O.fav (Orbicella faveolata, coral), and S.pur (Stronglyocentrotus purpuratus, sea urchin). Green asterisks mark channels we functionally compared. Sequences used in phylogeny construction are provided in Dataset S2. A scale bar indicates substitutions per site, and posterior probabilities <0.97 are given at branch nodes.

We functionally expressed Nematostella KCNQ (NvKCNQ) and compared it to human KCNQ2/KCNQ3 heteromers, a major M channel found in vivo in vertebrate nervous systems (30). NvKCNQ activates more slowly than HsKCNQ2/3 and has a flatter conductance–voltage curve (Fig. 8 A and B), but retains the low-threshold activation, which is believed to allow KCNQ channels to control excitation threshold. NvKCNQ reaches half-maximal activation (V₅₀) around −13.6 ± 1.5 mV (n = 14), significantly more positive than HsKCNQ2/3 channels (V₅₀ = −36.6 ± 1.6 mV; n = 10), but activation of both channels can be observed at voltages below −40 mV (Fig. 8 A and B). We coexpressed a voltage-sensitive phosphoinositide phosphatase from Ciona intestinalis, VSP, with NvKCNQ to test PIP₂ dependence. VSP dephosphorylates PIP₂ into PI₄P when activated by membrane depolarization, and can reduce PIP₂ levels in the oocyte membrane (67). If NvKCNQ requires PIP₂ for activation, a decrease in current should be observed during repeated depolarization in the presence of VSP. NvKCNQ currents reduced in magnitude during repeated steps to 0 mV from a holding potential of −100 mV when coexpressed with VSP, but not in controls (Fig. 8 C and D). HsKCNQ2/3 currents were similarly reduced in repeated 2-s steps to +40 mV specifically when coexpressed with VSP (Fig. 8 E and F). We did not observe a significant effect of VSP on HsKCNQ2/3 at 0 mV (where less VSP activation occurs), and a single sweep to +40 mV was sufficient to eliminate NvKCNQ currents. Both results suggest NvKCNQ has a lower affinity for PIP₂, and a smaller decrease in PIP₂ abundance may be sufficient to eliminate activation. Coexpression of NvKCNQ with PI(4)P5-kinase (PIP5K), which catalyzes PIP₂ production from PI₄P (68), increased peak NvKCNQ current size during 2-s voltage step to 40 mV (1 d postinjection) from 0.43 ± 0.09 μA (n = 8) in controls to 0.98 ± 0.16 μA (n = 10). This indicates that NvKCNQ is not PIP₂ saturated at resting Xenopus oocyte PIP₂ concentration. The VSP and PIP5K coexpression experiments together demonstrate PIP₂ dependence for NvKCNQ. Thus, PIP₂-dependent gating of KCNQ channels evolved before the cnidarian/bilaterian divergence.

Fig. 8.

Nematostella KCNQ encodes a subthreshold PIP₂-sensitive voltage-gated K⁺ channel. (A) Current traces from oocytes expressing Nematostella (NvKCNQ, Left) or human (HsKCNQ2/3, Right) KCNQ family channels. Currents were elicited by 2-s voltage steps from −120 to 60 mV in 20-mV increments (holding at −100 mV; tails at −50 mV). HsKCNQ2 and HsKCNQ3 were injected at a 1:1 ratio. (B) Normalized GV curves for NvKCNQ and HsKCNQ2/3 measured from isochronal tail currents at −50 mV following 2-s voltage steps to the indicated voltages. Curves show fits to a single-Boltzmann distribution; V₅₀ and slope values are reported in Table 1. (C) Example NvKCNQ current traces recorded in response to a series of 15 2-s voltage sweeps to 0 mV for NvKCNQ expressed alone (Left) or coexpressed with the Ciona VSP to progressively deplete PIP₂ (Right). (D) Peak current sizes observed in sweep 3 and sweep 10 relative to the peak current sizes of the first sweep. Error bars show SEM; n = 9 (control), 11 (VSP). The triple asterisk (***) denotes significance at P < 0.001 (t test). (E and F) Experiments were repeated for HsKCNQ2/3; voltage steps were taken to 40 mV, n = 10 (control) and 15 (VSP).

The structural basis of PIP₂ sensitivity in KCNQ channels has been studied using mammalian channels. Two separate binding sites for PIP₂ have been proposed: one that couples the voltage sensor to the activation gate and includes positively charged residues in the intracellular linkers of the voltage sensor and near the S6 activation gate of the pore (35), and one that implicates positively charged residues in the distal C terminus (69). Sequence alignments of 13 KCNQ sequences from eight species, including NvKCNQ, show high conservation of putative PIP₂-binding residues of the voltage sensor/activation gate site (Fig. S5). In contrast, we observed no significant alignment of the proposed C terminus binding site between KCNQ channels from distant species. We therefore suggest that the major evolutionarily conserved site for PIP₂ modulation of KCNQ channels resides at the voltage-sensor/activation gate interface. It is possible that the proposed C terminus binding site found in some vertebrate KCNQ channels (69) may contribute to their high PIP₂ affinity.

Discussion

Our results combined with previous studies (15, 16, 23, 29, 55) show that the functional and molecular diversification of Shaker and KCNQ families of voltage-gated K⁺ channels was largely complete before the divergence of cnidarians and bilaterians. Furthermore, Erg K⁺ channels, which constitute one of three bilaterian Ether-a-go-go gene subfamilies, are also highly conserved on functional level between cnidarians and vertebrates (70). The other two Ether-a-go-go subfamilies, Elk and Eag, have been identified in Nematostella (1). Thus, eight major classes of voltage-gated K⁺ channel are conserved between cnidarians and bilaterians, and the characteristic functional properties of six (Shaker, Shab, Shal, Shaw, KCNQ, and Erg) have now been shown to have evolved before the cnidarian/bilaterian divergence. Further support for an early diversification of voltage-gated K⁺ channels within parahoxozoans is the fact that Shaker, Shab, Shaw, and Erg channels can also be found in the placozoan Trichoplax adhaerens (65). Bilaterians are unique only in sharing two distinct KCNQ lineages; both KCNQ channel types are PIP₂ dependent (34, 40), and lineage-specific functional properties have not yet been described. Thus, ancestral parahoxozoans had a set of functionally diverse voltage-gated K⁺ channels that was sufficiently adaptable to regulate excitability in the highly diverse nervous systems of extant cnidarians and bilaterians. The evolution of voltage-gated Ca²⁺ channels tells a similar story; parahoxozoans share L-type, T-type, and N/P/Q/R-type channels, but ctenophores only have an N/P/Q/R-type channel (71).

Nematostella has single Shab and KCNQ genes, but contains large and functionally diverse expansions of the Shaker, Shal, and Shaw subfamilies. Interestingly, many of the genes in these expansions encode regulatory subunits that only function in heteromeric channels. The large Shab family expansion in vertebrates also contains regulatory subunits (58, 59). The regulatory subunit phenotype has therefore evolved independently in all four gene subfamilies of the Shaker family during metazoan evolution. These gene expansions provide increased functional diversity to specific subfamilies within various metazoan phylogenetic groups. Nevertheless, each of the voltage-gated K⁺ channel types we have examined has characteristic functional properties that have been highly conserved throughout parahoxozoan evolution. These include the delayed rectifier phenotype of Shab channels, the A-current phenotype of Shal channels, the rapid activation and steep voltage dependence of Shaker channels (16), the low activation threshold and PIP₂ dependence of KCNQ channels, and the I_Kr phenotype of Erg channels (59). The ancestral properties of Shaw channels are less clear, but our studies show a high degree of similarity between ecdysozoan and cnidarian Shaw currents. The steeply voltage-dependent and rapidly activating high-threshold Shaw currents of vertebrates may therefore represent a recent adaption specialized for rapid firing (24, 25).

Our results also demonstrate that many of the voltage-gated K⁺ channels that regulate excitability in bilaterian nervous systems are absent in the basal metazoan ancestor of ctenophores, sponges, and parahoxozoans. Shab, Shal, Shaw, and KCNQ channels are all found exclusively within the parahoxozoan lineage. Shaker subfamily channels, in contrast, are older and may have been present in a common ancestor of ctenophores and parahoxozoans, as well as at least one sponge lineage based on transcriptome sequences. We show evidence that two Mnemiopsis Shaker family channels from a clade that is closely associated with the parahoxozoan Shaker subfamily in phylogenies coassemble with Shaker subfamily channels from Nematostella and mouse. The ability for subunits to coassemble is considered a defining property of gene subfamilies (72) within the Shaker family. Our results therefore suggest that the Shaker subfamily emerged before the ctenophore/parahoxozoan divergence. Functional characterization of pure Mnemiopsis Shaker channels, which we were unable to obtain in this study, would resolve whether there is close conservation of biophysical properties between the ctenophore and parahoxozoan Shaker subfamilies.

The high conservation of functional phenotype within K⁺ gene families over long evolutionary distances suggests that at least some important aspects of the parahoxozoan voltage-gated K⁺ current diversity will be missing in ctenophores. However, our results do not necessarily mean that the complexity of electrical signaling is “simple” in the ctenophore nervous systems. Mnemiopsis leidyi has 49 Shaker family genes, more than we have observed in any other species, and these fall into two distinct clades. One possibility is that the clades represent functionally distinct gene subfamilies. However, because we have so far been unable to express ctenophore channels, we do not yet know the extent of their functional diversity. It is entirely possible that some of the diversity provided by parahoxozoan-specific Shaker K⁺ channel subfamilies could be provided by functional diversification within the large Mnemiopsis gene expansions. Future cross-species comparisons of ctenophore Shaker family channels will be needed to determine how the diversity we observed in Mnemiopsis evolved within the ctenophore lineage. What is clear from our findings is that if ctenophore voltage-gated K⁺ channels are functionally diverse, that diversity evolved separately within ctenophores.

The selective pressure that drove evolutionary diversification of voltage-gated K⁺ channels in ancestral parahoxozoans has not yet been determined. If the hypothesis that the nervous system evolved separately in ctenophores and parahoxozoans is true (42, 45), then it may have been the de novo evolution of neurons and muscle within the parahoxozoan lineage that provided a selective pressure for the diversification of electrical signaling mechanisms. However, if nervous systems were present in the ctenophore/parahoxozoan ancestor, what might have driven the evolution of so many new channel types in parahoxozoans? We can only speculate because the evolution of nervous system structure and function is still poorly understood. Parahoxozoan-specific channels (Shab, Shal, Shaw, and KCNQ) specifically regulate firing threshold or complex action potential patterning, whereas Shaker channels, which our study suggests are shared by ctenophores and parahoxozoans, play a critical role in axonal action potential propagation. It is therefore tempting to speculate that voltage-gated K⁺ channel evolution in basal metazoans was driven by the evolution of action potential propagation, whereas parahoxozoan-specific K⁺ channel diversification accompanied more complex patterning of signaling within networks. However, further characterization of cnidarian and ctenophore channels, neurons, and circuits on functional level will be needed to better understand how the evolution of voltage-gated K⁺ channels fits into the broader picture of the evolution of neuronal function.

Methods

Gene Identification, Cloning, and Phylogenetic Analysis.

Cnidarian and Mnemiopsis Shaker and KCNQ family K⁺ channel genes described in this study were identified and compiled through comprehensive BLAST (46) searches of genome drafts, transcriptomes, and gene predictions of Mnemiopsis leidyi (41) and Nematostella vectensis (73), and for KCNQ only, Hydra magnipapillata (66), and Acropora digitifera (74) and Orbicella faveolata. Multiple bilaterian members of each channel type were used as query sequences, and reciprocal BLAST searches of identified sequences against bilaterian databases were used to classify the sequences before phylogenetic analysis. Most queries identified all voltage-gated K⁺, Na⁺, and Ca²⁺ channels, but reciprocal searches sorted target sequences by gene family and were used to refine gene predictions when necessary.

Alignments for phylogenetic analysis were produced using the Muscle Algorithm as implemented in MEGA 6 (75), and regions of low length conservation were trimmed before phylogenetic analysis. Shaker family alignments included the N-terminal T1 domain and S1–S6 voltage-gated K⁺ channel core. KCNQ alignments contained the channel core and KCNQ-specific coiled-coil region of the C terminus. Sequences used for phylogenetic analysis are provided in Dataset S1 (Shaker) and Dataset S2 (KCNQ). Bayesian inference phylogenies of channel proteins were constructed using MrBayes (76). Analyses were run under a mixed model for 1 million generations with two runs of four chains. The first 25% of the analysis was discarded as a burn in phase, and the displayed phylogenies are the consensus. The convergence diagnostic potential scale reduction factor was 1.000 for Shaker and 0.999 for KCNQ, indicating appropriate convergence of independent runs. Minimum-evolution, maximum-likelihood, and SSN analyses are described in SI Methods.

Nematostella and Mnemiopsis channel genes were cloned using standard RT-PCR techniques from total RNA samples from whole adult animals. Briefly, 500 ng to 2 μg of total RNA was reverse transcribed using oligo-dT, and gene-specific primers were used to amplify coding regions. When necessary, overlap extension PCR was used to assemble full-length coding regions from overlapping fragments. Coding regions were cloned into the pOX vector (23) for expression in Xenopus oocytes and sequence confirmed. Consensus DNA coding sequences are reported in Dataset S3, and only clones matching these sequences were used for expression analysis.

Electrophysiology.

Xenopus laevis ovaries were obtained from Xenopus I. Mature oocytes were enzymatically defolliculated with type II collagenase (Sigma-Aldrich) at 1 mg/mL in Ca²⁺-free ND98 solution (98 mM NaCl, 2 mM KCl, 1 mM MgCl₂, 5 mM Hepes, pH 7.2). Following digestion, oocytes were maintained in ND98 culture solution (98 mM NaCl, 2 mM KCl, 1.8 mM CaCl₂, 1 mM MgCl₂, 2.5 mM Na-pyruvate, 100 U/mL penicillin, 100 µg/mL streptomycin, and 5 mM Hepes, pH 7.2). Capped, polyadenylated cRNA transcripts were generated from linearized expression plasmids using T3 mMessage mMachine kit and Poly-A tailing kit (Life Technologies). LiCl precipitation was used to purify transcripts, and pellets were resuspended in nuclease-free water supplemented with RNase inhibitor. Transcripts were injected in 50-nL volumes into mature oocytes, which were incubated at 18 °C in ND98 culture solution for 1–3 d before recording.

Two-electrode voltage-clamp recordings were carried out at room temperature (22–24 °C) using a Dagan CA-1B amplifier, and data were collected and analyzed with the pClamp 10 acquisition suite (Molecular Devices). Recordings were performed under constant perfusion (98 mM NaOH, 2 mM KCl, 1 mM CaCl₂, 1 mM MgCl₂, and 5 mM Hepes, pH 7.0). Electrodes (0.4–1 MΩ) were filled with 3 M KCl and bath-clamp circuitry was isolated with a 1 M NaCl/agarose bridge. Currents were sampled at 10 kHz and filtered at 2 kHz. Hardware capacitance and leak compensation was used in some recordings.

Data were fit and plotted in Origin 8.1 (OriginLab). Single-Boltzmann distributions were fit using the equation f(V) = A₂ + (A₁ − A₂)/(1+e^(V−V50)/s), where V₅₀ is the half-maximal activation voltage, s is the slope factor, and A₁ and A₂ are the lower and upper asymptotes, respectively. Double-Boltzmann distributions were fit using the equation f(V) = y₀ + A[p/(1 + e^(v−x01)/s1) + (1 − p)/(1 + e^(v−x02)/s2)], where y₀ is the current offset, A is the amplitude span, and p is the fraction of current component in the more hyperpolarized voltage range. Data points show averages of normalized values for individual cells. Arithmetic means of V₅₀ and s values from individual measurements were used to generate the Boltzmann fits shown in figures. Recovery time course for NvShal channels was fit with two exponentials using the equation f(t) = A₁*exp(−x/t₁) + A₂*exp(−x/t₂) + y₀ (x, time interval of recovery; t₁ and t₂, time constants of recovery; A₁ and A₂, amplitudes of the recovering components; y₀, current offset). Peak current amplitudes measured from test pulses were normalized to the peak current amplitude of the first pulse. Data points are averages of measurements from individual cells, and fits shown were generated with arithmetic means of fit parameters.

Statistical comparisons between datasets were carried out using two-tailed t test.