Contractions in 12ppt ASW. N. vectensis polyps occasionally undergo a slow peristaltic contraction that starts about halfway down the body column and ends at the foot or physa (Fig. 1a; Supplementary Video 1). The waves travel at 9.7 ± 2.0 µm/s (6 animals, 17 measurements) (Fig. 1b). In no case did we observe a second wave begin in an animal before the first wave had ended. Occasionally, we observed a wave traveling in the reverse direction (physa to mid-column).
Polyps underwent an average of 2.5 peristaltic waves per 5 min interval (pooled measurements from 808 animals that were subsequently used in experiments with different treatments). However, the number of waves varied from zero to nine with a median value of 2 and was not normally distributed (Fig. 2a). We fitted the distribution to the sum of two weighted Poisson distributions. The means and 95% confidence limits of the fit were 0.47 ± 0.11 waves per 5 min (0.41 ± 0.05 weight) and 3.94 ± 0.34 waves per 5 min. This suggests that N. vectensis polyps normally have two activity levels: low and high, with slightly more than half of the polyps being in the high activity group.
Wave frequency is increased by nicotine. The addition of nicotine (final concentration 185µM) to the solution bathing the polyps dramatically increased the number of waves to an average of 6.0 waves per 5 min (median of 6 waves per 5 min, 187 animals, Supplementary Video 2). We fitted the distribution as the sum of two Poisson distributions but the fitted weight of one component was zero; the remaining Poisson component had a mean of 6.5 ± 0.83 waves per 5 min (Fig. 2b). Thus, all of the polyps contracted at an activity level higher than that of the high activity in the control. The speed of the contractions in the presence of 185 µM nicotine was twice as fast as those under control conditions (Fig. 2c; 19.3 ± 3.7 µm/s, 5 animals, 21 measurements, p < 0.001, t-test, the same set of animals as were used to obtain the control speed, Supplementary Figure S1).
The excitation induced when nicotine is added to the well is not due to the animals being disturbed by the addition of fluid. Figure 3 shows the results for a set of 56 animals recorded under control conditions and then again five minutes after the addition of 0.10 ml of 12 ppt ASW. There was no significant change in the distribution of waves. These same animals responded to the subsequent addition of nicotine to the well (final concentration 370 µM).
The stimulating effect of nicotine was dose dependent (Fig. 4). For three concentrations of nicotine, 93 µM (63 animals), 185 µM (169 animals) and 370 µM (55 animals) and the corresponding controls, the mean frequency of waves increased by factors of 1.8, 2.4 and 4.3, respectively. Linear regression of the waves vs. nicotine concentration data yielded a slope of 0.015 ± 0.002 waves/µM (95% confidence limits) indicating that the dose-dependence is highly significant (Supplementary Figure S2).
Contractions in the presence of acetylcholine (ACh). Surprisingly, ACh did not stimulate peristalsis. At concentrations ranging from 10 µM to 5 mM, ACh appeared to inhibit peristalsis, but the results were inconsistent. In the example shown in Fig. 5a, 500 µM ACh had no effect on the frequency of waves. One possible explanation for the lack of a stimulatory effect of ACh is that acetylcholinesterases in N. vectensis may rapidly hydrolyze exogenously applied ACh. We tested this idea by first exposing polyps to the acetylcholinesterase inhibitor, edrophonium and then added ACh. 500 µM edrophonium increased the frequency of waves from 1.50 per 5 min to 3.32 per 5 min (Fig. 5b). Subsequent addition of 500 µM ACh increased the frequency of waves even further to 5.21 per 5 min. This suggests that inhibition of acetylcholinesterase by 500 µM edrophonium increases the effectiveness of both endogenous and exogenously applied ACh.
N. vectensiscontains a large number of nAChR paralogs. We hypothesized that our behavioral observation of dramatic, dose-dependent peristalsis in response to nicotine is due to the presence of nAChRs. To test this hypothesis, we first identified all potential nAChR paralogs in the N. vectensis genome. To determine the extent that the details of this type of neurochemical signaling may be homologous or convergent with bilaterian systems, we determined the phylogenetic relationship of the N. vectensis nAChRs to other characterized nAChR genes. According to our conservative estimate, the N. vectensis proteome contains 49 nAChR copies (Supplementary Table S3), the most of any cnidarian sampled to date. All of these encoded proteins have the characteristic structural features found in mammalian nAChRs (Supplementary Figure S3) and many of the amino acid residues within the agonist/antagonist binding loops are identical to those found in human nAChR proteins (Supplementary Table S4). These paralogous proteins are encoded on 27 different scaffolds. Queries into previous expression sets indicate that all 49 genes examined are expressed with varying temporal dynamics and magnitude 44–46. The 15 species of cnidarians we analyzed average 34 nAChR paralogs, ranging from 13 in H. vulgaris to 46 in Pocillopora damicornis (Supplementary Table S2). The sampled bilaterian proteomes contain, on average, 31 nAChR paralogs (including Zinc/5HT3 receptors), ranging from 7 in Ciona intestinalis (ascidian) to 68 in Mizuhopecten yessoensis (scallop). Humans have 16 nAChR paralogs. We verified that the number of predicted nAChR genes in a genome is not strongly predicted by the completeness of the genome annotation (Supplementary Figure S4). In total, our final protein alignment includes 502 cnidarian genes and 498 bilaterian genes and contains 435 positions (Supplementary Table S5).
A single lineage of nAChR genes underwent a dramatic expansion in Cnidaria. Our phylogeny is rooted with a zinc/5HT3 outgroup, the paralogous sister group to all other cationic subunits (“Gene Tree” in Fig. 6; Supplementary Figure S5). In this tree, the relationships among the bilaterian nAChRs generally agrees with typical relationships described in recent literature (e.g.47). Within this phylogeny, we observe the 502 cnidarian gene copies occur in two distinct clades. The first cnidarian clade contains genes present in a single copy in just five of the anthozoans (Fungia spp., Stylophora pistillata, Pocillopora damicornis, Acropora tenuis, and Porites asteroides). These five genes (“Cnidaria-2” in Fig. 6) are part of the divergent group (Branch E) related to Drosophila β3 and found in xenoceolemorphs, protostomes, echinoderms, and hemichordates, but not in chordates.
The remaining 497 cnidarian genes (99%) belong in a single large, cnidarian-specific clade; we call this clade the Cnidaria-group of nAChRs (“Cnidaria-group”) (* in Fig. 6). All 49 N. vectensis nAChRs are found in the Cnidaria-group (purple polyps in Fig. 6). The Cnidaria-group is well supported as a clade (98.9% SH-aLRT, 100% UFboot support). The Cnidaria-group diverged after the α9/10 lineage and its relatives (branch D ; found in protostomes, echinoderms, hemichordates, and chordates). The Cnidaria-group is sister to a clade that includes the “B1-group” and the α7 group (branch A in Fig. 6) (67.2% SH-aLRT, 60% UFboot support). The B1-group is a large clade of well-studied bilaterian nAChRs, including the α1–6, β1–4, δ, ε, and γ subunits (branch B; found in all bilaterians studied, including the xenoceolemorph). The α7 group is found in protostomes, echinoderms, hemichordates, and chordates (branch C).
We tested our confidence in the placement of the Cnidaria-group as sister (closest relatives) to the B1 + α7 group using alternative topology tests (Supplementary Table S6), and were able to reject alternative placement (c-ELW and p-AU < 0.01 for both; joining along branches B or C in Fig. 6). All other sister relationships to major groups we tested cannot be rejected by either test (p > 0.05). We cannot reject the Cnidaria-group as sister to the α9/10 group (branch D; found in protostomes, echinoderms, hemichordates, and chordates), the Dβ3/divergent group 48 (branch E; found in protostomes, echinoderms, hemichordates, and some cnidarians; see below), B1 + α7 + α9/10 (branch F), or B1 + α7 + α9/10 + Dβ3 (branch G).
It appears that the 497 gene copies in the Cnidaria-group all expanded from a single common ancestral gene in ancient cnidarians. Gene duplications led to increases in nAChR copy number especially in anthozoans, including the Hexacorallia, and Scleractinia cndiarians. Intriguingly, this expansion in cnidarians tentatively appears to be the result of an expansion of the same stem lineage that also exhibited a large expansion in bilaterian lineages (B1-group; although we cannot reject some alternatives). This brings about several important questions. First, why was this specific lineage subject to dramatic expansion, independently but in parallel (convergently), in bilaterians and cnidarians? Second, in how many ways did the nAChR systems in bilaterians and cnidarians evolve in parallel? Here, we have shown that they evolved in parallel in terms of expansion of the gene family copy number, suggesting the convergent evolution of a complex nAChR system. However, do the same types of subunits exist? Are they assembled into homopentamers and heteropentamers of similar types to those of bilaterians? Are they equivalently tissue specific? And do the serve the same range of functions, or do they have new and marvelous functions to discover? Answers to these questions will require further phylogenetic and functional characterization.
The observation that various types of complexity has arisen in animals multiple times, and is not specific to bilaterians, is now becoming well-appreciated 49. Indeed, the independent expansion of nAChR gene family copy number in bilaterians and cnidarians that we observe here is consistent with a similar inference for the entire ligand-gated, cys-loop receptor superfamily (which also includes GABAA and Glycine anionic receptors; not considered here) in a study using two cnidarian genomes 50. That study also inferred a similar pattern for glutamate-gated channels and voltage gated potassium channels, also suggesting repeated and independent occurrences of the expansion of nervous system complexity.
To consider the question of how early nAChRs evolved, we also searched in proteomes from earlier diverging metazoan lineages. We found no evidence of nAChRs in the poriferans Xestospongia testudinaria, Tethya wilhelma, Stylissa carteri, Ephydatia muelleri, Stylissa carteri, and Amphimedon queenslandica, or in the ctenophores Mnemiopsis leidyi and Pleurobrachia bachei (not shown). Accumulating phylogenomic evidence suggests that Bilateria and Cnidaria form a monophyletic group that does not include the Porifera and Ctenophora (e.g. 51–53; although other work places Porifera as sister to cnidarians, as in 54. The presence of nAChRs in Bilateria and Cnidaria supports their sister relationship and monophyly, with respect to poriferans and ctenophores; this distribution is the same as for the entire cys-loop receptor superfamily as well as acid sensing channel 50, and has also been observed for other types as genes, such as for Hox/ParaHox, S50 and K50 PRD and HNF class homeodomains 55.
Repeated evolution of non-αsubunits. Of the 49 nAChR genes in N. vectensis, 44 encode α-like subunits (90%), as defined by the presence of a double-cysteine in loop C (Supplementary Figure S3). The five genes encoding non-α subunits come from four clades that each evolved independently into the non-α state (Fig. 6, Supplementary Figure S5). Among all cnidarian nAChR genes, 83% (417/502) are α-like. In contrast, for bilaterians, only 57% are α-like (256/453). In both cases, the non-α state has arisen independently numerous times from the ancestral, α-like nAChRs ancestor. Like Dβ3, the Cnidaria-2 group is non-α.
There are no muscarinic cholinergic receptors inN. vectensis. To rule out the possibility that the behavioral, dose dependent peristalsis we observed is due to the presence of muscarinic cholinergic receptors (mAChRs), we searched for the closest related genes to mAChRs within N. vectensis and determined their phylogenetic placement among rhodopsin-like G-protein-coupled receptors. Previous work has shown that anthozoans like N. vectensis do not possess mAChRs (Anctil, 2009; Faltine-Gonzalez and Layden, 2019) whereas hydrozoans such as H. vulgaris do (Collins, 2013.) We therefore conducted a phylogenetic analysis of mAChRs and other related rhodopsin-like G-protein-coupled muscarinic receptors. We identified the 10 top matching genes to a mAChR-specific HMM from each H. vulgaris and N. vectensis (Supplementary Figure S6). We found genes from H. vulgaris are sister to the bilaterian mAChRs. In contrast, the N. vectensis genes are related to octopamine receptors, rather than mAChRs, consistent with a lack of mAChRs. We also compared these cnidarian sequences at the 14 amino acid sites known to be critical to ACh binding (Supplementary Table S7). We found that the H. vulgaris genes have an average of 8 identical amino acids (range 7–14) while N. vectensis genes have an average of only 3 critical amino acids (range 2–5). We therefore infer that the response we see in N. vectensis is mediated by nAChRs, rather than mAChRs.