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Peripheral and central employment of acid-sensing ion channels during early bilaterian evolution

Acid-sensing ion channels (ASICs) are membrane proteins that endow vertebrate neurons with fast excitatory responses to decreases in extracellular pH. Although previous studies suggest that ASICs are found in certain invertebrates, the lineage in which ASICs emerged and the functional role of ASICs beyond the vertebrates is unknown. We reconstructed ASIC evolution by surveying metazoan ASICs and performing a phylogenetic analysis, which suggests that ASICs evolved in an early bilaterian. This was supported by electrophysiological measurements of proton-gated currents at heterologous channels from diverse bilaterians. This also revealed substantial variation in biophysical properties of broadly related ASICs, with selective sodium/potassium ion permeability ranging from 3-to 36-fold and half-maximal activating pH from 4.2 to 8.1. Furthermore, we studied the expression of ASICs in species outside the vertebrates and found expression in the central nervous system of certain bilaterian lineages but in the periphery, including nervous system, digestive system, and motile ciliated epithelia, of most lineages. The loss of such epithelial cells in Ecdysozoa might explain the loss of ASICs from this major animal lineage. Our results suggest that ASICs emerged in an early bilaterian, and expression in the central nervous system was accompanied or even pre-dated by expression in the periphery.


Introduction
Much of the morphological and behavioral complexity of animals is enabled by nervous systems in which neurons in the periphery sense and convey information from the environment, and more central neurons integrate and dispatch information to effectors [1,2]. Such rapid sensory (environment → cell) and synaptic (cell → cell) signals rely on ligand-gated ion channels (LGICs), membrane proteins that convert chemical messages into ionic currents within milliseconds [3,4]. These channels are found in all multicellular animals (Metazoa) and even in outgroup lineages such as bacteria and plants [5][6][7]. However, animals with nervous systems (Ctenophora, Cnidaria, and Bilateria) present a larger and more diverse LGIC gene content -i.e., more Cys-loop receptors, ionotropic glutamate receptors, and degenerin/epithelial sodium channel (DEG/ENaC) genes -than animals without nervous systems (Porifera and Placozoa) [8]. Although the evolution of the original nervous system(s) did not necessarily involve a rapid expansion of LGICs, the elaboration and refinement of nervous systems within particular lineages did, and this likely endowed bilaterian neurons with a sophisticated chemo-electrical toolbox in the ancestors of today's complex bilaterian animals [1,8,9]. Unfortunately, studies addressing the functional contribution of LGICs to early bilaterians are lacking [10].
Within the DEG/ENaC family of voltage-insensitive sodium channels, several independent expansions of neuronally expressed subfamilies have occurred, including degenerin channels (DEGs) in nematodes, pickpocket channels (PPKs) in arthropods, peptide-gated channels (HyNaCs) in cnidarians, and acidsensing ion channels (ASICs) in vertebrates [11][12][13][14][15] (Fig 1A). ASICs are proton-gated ion channels that in rodents are widely expressed in the nervous system, with scattered expression in other cells, such as epithelia [16] (Fig 1B,C). In rodents and humans, ASICs are expressed or stimulus-upregulated in sensory neurons innervating skin, joints, and the gastrointestinal tract where their activation by acid or other inflammatory stimuli mediates touch or causes pain or hyperalgesia [17][18][19][20], typifying a chemosensory role. More recent experiments in rodents show that ASICs expressed postsynaptically in central neurons mediate depolarization in response to brief drops in synaptic pH during neurotransmission [21,22], and ASICs thus perform an additional central role, mediating excitatory signals between neurons. Such roles might extend beyond the vertebrates, however, as it was recently discovered that functional ASICs exist in all Deuterostomia members (tunicates, cephalochordates, hemichordates and echinoderms), indicating that the ASIC subfamily is older than previously thought [12]. As ASICs are expressed in both central and peripheral neurons of tunicates [12,23] and in sensory neurons of echinoderms [24] (Fig 1B), it is difficult to assess the functional role of ASICs in early deuterostomes before they diversified.
It also remains unclear whether the ASIC subfamily might have emerged much earlier than the first deuterostomes and subsequent centralization of their nervous system, and what original function ASICs might have fulfilled earlier in metazoan evolution. Resolving this would require better resolution of the phylogenetic origin of ASICs. Furthermore, investigation of the expression of ASICs in a broader selection of species is required to address the evolution of peripheral, sensory and central, synaptic roles. We therefore performed a thorough phylogenetic investigation of metazoan DEG/ENaC genes, with a focus on ASICs, using unexplored transcriptomes and genomes. Moreover, we carried out expression analysis by in situ hybridization and determined the electrophysiological properties of several diverse ASICs from phylogenetically informative species. Results from these experiments enable a new hypothesis on the evolution and function of ASICs.  [11,25]). (B) Phylogenetic tree of main animal groups highlighting known ASIC expression (blue) and previously suggested ASIC emergence (blue star). Dotted lines show alternative positions for Ctenophora and Xenacoelomorpha. Cartoons show previously described ASIC protein and/or RNA expression in Chordata (rodent brain, spinal cord, lung, heart, kidney, gut, bladder, testis, bone, eye, ear, and skin [26]; tunicate larva sensory vesicle and bipolar tail neurons [23]) and Ambulacraria (echinoderm larva ciliary band [24]). (C) ASIC function. At rest, the channel is closed and impermeable. Upon extracellular acidification, certain amino acid side chains in the ASIC extracellular domain are protonated, causing conformational changes that open the channel, allowing sodium ions to flow down their chemo-electric gradient across the membrane. Genes analyzed experimentally in this study are indicated by † (gene expression) and/or * (electrophysiology). Scale bar, amino acid substitutions per site. aLRT SH-like (black) and aBayes (magenta) likelihood-based support values indicated.

ASICs are expressed in two domains in Xenacoelomorpha
These results suggest that ASICs emerged during early bilaterian evolution, and we next sought evidence for potential biological roles by investigating the expression of ASIC genes in the different bilaterian lineages. The precise relationships between these lineages are under debate, but Xenacoelomorpha forms a putative sister group to all remaining Bilateria [27][28][29][30]. Therefore, we investigated ASIC expression in Xenacoelomorpha, utilizing the acoels Isodiametra pulchra, Hofstenia miamia, and Convolutriloba macropyga. Xenacoelomorphs show a large variation in nervous system architecture, but all possess a basal epidermal nerve plexus and some species, more internally, an anterior condensation of neurons or "brain" [31]. I. pulchra and C. macropyga also possess longitudinal bundles of neurons [32][33][34]. External to this nervous system in acoels is generally a sheet of longitudinal and ring muscles and, more externally, ciliated epithelial cells mediating locomotion [35][36][37].
From a dorsal view, expression of I. pulchra ASIC mRNA can be detected in several cells across the central anterior of the adult animal ( Fig 3A). This is the location of the I. pulchra brain, where e.g. serotonergic and peptidergic neurons form lateral lobes, connected by a frontal ring and a posterior commissure, and from which four pairs of nerve cords extend posteriorly ( Fig 3A [32]). Fluorescent confocal micrographs show that the ASIC-expressing cells are associated with the posterior commissure of the brain and the dorsal and lateral neurite bundles (Fig 3A, white arrowheads). H. miamia possesses an external body wall muscle, and more internally, tissue Layer I, another muscle wall, and then tissue Layer II [38]. There is a relatively loose anterior condensation of e.g. GABAergic, dopaminergic, and peptidergic neurons in the anterior third of the animal in layer II, and various peripheral neurons are found throughout the animal in both layers (Fig 3B, [38]). In the young adult H. miamia, we observed high ASIC expression in cells throughout the anterior condensation ( Fig 3B). Similar to I. pulchra, the ASIC signal is strongest near the dorsal commissure but, in contrast, also in scattered cells across the animal, including in the very periphery ( Fig 3B, yellow arrowhead). The latter are present in inner and outer layers and are thus consistent with the dispersed, diverse neurons found in layers I and II of the animal. C. macropyga ASIC expression was high and broad. In the anterior, ASIC-expressing cells are clearly associated with the brain (Fig 3C', white arrowheads). Throughout the animal, including the very periphery, there is medium to high ASIC expression. While perhaps indicative of expression in locomotory ciliated cells covering the animal, this expression may be part of the extensive nerve plexus (Fig 3C''). In summary, I. pulchra shows localized, central ASIC expression whereas C. macropyga and H. miamia show both central and scattered, peripheral ASIC expression. Thus, ASIC is expressed in two distinct domains in Xenacoelomorpha: a central domain, and a peripheral domain, and ASICs are likely to have performed central, integrative and peripheral, sensory roles in early acoels. after [33]. ant, anterior; dor, dorsal; nb, neurite bundles; pc, posterior commissure. Mid-right: protongated currents in Xenopus laevis oocytes expressing xenacoelomorph ASICs (scale bars: x, 5 s; y, 0.5 μA). Lower right: Mean (± SEM) pH 4-gated current (I, μA) at different membrane potentials (V, mV) in the presence of 96 mM extracellular NaCl or KCl (n = 7-8). Reversal potential (V rev ) was read off these plots and the difference between V rev,NaCl and V rev,KCl was used to calculate relative ion permeability (P Na+ /P K+ , Materials and Methods). (C) ASIC mRNA expression (magenta) and tyrosinated tubulin immunoreactivity in C. macropyga. As C. macropyga is larger, images of slightly ventral (C') and dorsal (C'') planes are shown. (D) Filled symbols: mean (± SEM) normalized current amplitude in response to increasing proton concentrations (activation, n = 6-10). Open symbols: mean (± SEM) normalized current amplitude in response to pH 4 following pre-incubation in decreasing pH (steady state desensitization, n = 4-6).

Xenacoelomorph ASICs mediate excitatory currents gated by high proton concentrations
When ASICs are exposed to drops in extracellular pH they typically show a transient depolarizing current (inward flow of Na + ions), rapidly followed by either desensitization-a non-ion-conducting state in the presence of agonist-or a smaller sustained current [39]. To test the function of xenacoelomorph ASICs, we injected the ASIC cRNAs into Xenopus laevis oocytes and measured membrane current in response to decreasing pH using two-electrode voltage clamp. At oocytes expressing I. pulchra ASIC, drops to pH 5.0 and lower rapidly activated a transient current (Fig 3A, mid-right). However, compared to other ASICs, responses to increased proton concentrations were relatively inconsistent at I. pulchra ASIC: although transient currents were never activated by pH higher than 5.6, the concentration dependence of the transient current between pH 5.3 and 4.0 was inconsistent, and a sustained current developed during prolonged proton application at pH ~4.0 in some cells and at pH ~3 in others ( Fig 3A, mid-right). Thus, I. pulchra ASIC is indeed gated by protons, but with relatively low potency (pH for half-maximal activation of transient current (pH 50 ) = 4.0 ± 0.2). The H. miamia ASIC also formed proton-gated channels, with rapid transient currents followed by smaller sustained currents in response to drops to pH 5.6 through 4.0 (pH 50 = 4.8 ± 0.1, Fig 3B,D). We observed no proton-gated currents in Xenopus oocytes injected with C. macropyga ASIC cRNA (n = 6), presumably due to low heterologous expression. Like vertebrate ASICs, I. pulchra and H. miamia ASICs showed decreased responses to activating pH (4.0) after preincubation in slightly decreased pH ( Fig 3D, open symbols), indicating that steady-state desensitization is a broadly conserved phenomenon.
Most ASICs so far studied have a slight preference for sodium over potassium ions, with a relative sodium/potassium ion permeability (P Na+ /P K+ ) of ~10, and thus activation of ASICs leads to depolarization and generation of action potentials in mice [40]. I. pulchra ASIC (P Na+ /P K+ = 8.3 ± 2.0) showed similar Na + selectivity to most ASICs ( Fig 3A, lower right), whereas H. miamia Na + selectivity (P Na+ /P K+ = 36 ± 11; Fig 3B) is remarkably high compared to most ASICs and is more reminiscent of the high preference for sodium seen in ENaCs [41]. Thus, both in xenacoelomorphs, which belong to the putative, sister group to all other bilaterians, and in deuterostomes, such as rodents and humans, ASICs are excitatory receptors for protons that are expressed highly in both the central nervous system and in peripheral neurons.

ASICs are expressed in peripheral neurons and the digestive system in Spiralia
Excitatory proton-gated currents and broad neuronal expression of ASICs in xenacoelomorphs is reminiscent of vertebrate ASICs. However, whether the ancestral bilaterian ASIC had this role remains unclear, as the position of Xenacoelomorpha as sister taxon of all other bilaterians or as part of a group with hemichordates and echinoderms within Deuterostomia is debated [27][28][29][30]. We therefore turned to the third major lineage of bilaterians and investigated previously unidentified ASICs in Protostomia. We investigated the expression and function of ASICs in brachiopod, phoronid, and annelid larvae, allowing us to study whole-animal gene expression in animals with typical spiralian features such as an anterior brain or apical organ and ventrolateral or peripherally extending nerve cords.
In the brachiopod Terebratalia transversa, ASIC was expressed most highly in cells at the lateral edge of the central neuropil in late larvae ( Fig 4A). Confocal images suggest that these ASIC-expressing neurons project latero-anteriorly ( Fig 4A, inset), suggesting an anterior sensory role, and these cells are slightly lateral/caudal to previously identified sensory neurons in the T. transversa apical organ [42], in close proximity to previously described cholinergic and peptidergic cells [33,43]. In the phoronid Phoronopsis harmeri, ASIC was expressed in two principal domains. Regarding the nervous system, there was a clear signal in relatively few cells in the perimeter of the hood (Fig 4B, black arrowheads). This area is innervated by the peripheral nerve ring and controls swimming by ciliary beating [44,45]. Whether the ASIC-positive cells have a role in chemosensing or controlling ciliary beating remains unclear. There was no ASIC expression in the more central parts of the nervous system, including both the apical organ, an anterior group of sensory neurons that gives way to the developing brain, and the main nerve ring that runs caudally from the apical organ through the trunk and innervates the tentacles [45][46][47]. The second domain, in which expression was even more prominent than that in the peripheral nervous system, was the digestive system. Here, expression was high in numerous cells around the digestive tract (comprising a stomach in the trunk and intestine in the pedicle at this stage), most noticeably just below the pharynx and around the lower half of the stomach (Fig 4B, orange arrowheads, [48]). We also detected such digestive system expression in the more distantly related spiralian, the annelid Owenia fusiformis. In dome-shaped O. fusiformis larvae 18-20 hours post fertilization (hpf), the young nervous system consists of a dorsal apical organ, several ventral prototroch (or ciliary band) neurons, and a small number of anterior and lateral neurons connecting these apical and peripheral structures [49,50]. In 19 hpf O. fusiformis, we observed no ASIC expression in the young nervous system. Instead, ASIC was expressed most clearly in cells around the oesophagus, and, more broadly, in the midgut (Fig 4C). These results show that ASICs are expressed in the peripheral nervous system and/or epithelial cells in the digestive system of all spiralians tested.  [50,51] and ASIC mRNA expression (ant, anterior; api, apical; lat, lateral; ven, ventral; scale bar, 40 µm). Arrowheads, ASIC expression; asterisks, unspecific staining common in T. transversa [33]. Lower left: protongated currents in Xenopus laevis oocytes expressing indicated spiralian ASICs (scale bars: x, 5 s; y, 1 μA). Lower right: mean (± SEM) pH 6.5-(L. anatina) or 4-(P. harmeri ASIC) gated current (I, μA) at different membrane potentials (V, mV) in the presence of 96 mM extracellular NaCl or KCl (n = 5-8). Reversal potential (Vrev) was read off these plots and the difference between V rev,NaCl and V rev,KCl was used to calculate relative ion permeability (P Na+ /P K+ ). (C) Left to right: Animal morphology, ASIC expression, proton-gated currents, and pH 6.0-gated current at different membrane potentials, as in (A,B) at O. fusiformis ASIC. (D) Filled symbols: mean (± SEM) normalized current amplitude in response to increasing proton concentrations (activation, "act.", n = 6-11). Open symbols: mean (± SEM) normalized current amplitude in response to pH 7 for O. fusiformis, 6.5 for L. anatina, and 4 for P. harmeri ASIC following pre-incubation in decreasing pH (steady state desensitization, n = 4-5).

Spiralian ASICs show a wide range of proton sensitivity and ion selectivity
We next tested the electrophysiological properties of spiralian ASICs by expressing them heterologously in Xenopus oocytes. We observed no proton-activated currents in Xenopus oocytes injected with T. transversa ASIC RNA (n = 10), and we cannot conclude if this is due to low heterologous expression or unknown function of this channel. However, we identified an ASIC transcript in another brachiopod, Lingula anatina [52], and observed functional expression of this channel. L. anatina ASIC was sensitive to relatively low proton concentrations, activated by pH in the range of 7.4 to 6.0 (Fig 4A, pH 50 = 7.3 ± 0.2) and showed typical selectivity for sodium over potassium ions (Fig 4A, P Na+ /P K+ = 9.0 ± 1.7). The phoronid P. harmeri ASIC showed much lower proton sensitivity, with small currents in response to pH 5.3 and lower that rapidly desensitized in the continued presence of low pH (Fig 4B, pH 50 = 5.1 ± 0.1) with almost no preference for sodium over potassium ions (Fig 4B, P Na+ /P K+ = 3.0 ± 0.9). We next tested the function of ASIC from the annelid O. fusiformis. This channel was also highly sensitive to low proton concentrations. When held at pH 7.5 and exposed to lower pH, O. fusiformis ASIC showed very small current responses, but when held at pH 9.0 and exposed to small drops in pH, large inward currents were activated that rapidly desensitized (Fig 4C,  pH 50  Spiralian ASICs seem to vary in apparent proton affinity and in ion selectivity among the lineages, from low proton sensitivity (pH ~5) and essentially non-selective cation currents in phoronid ASIC to high proton sensitivity (pH 7-8) and ~10-fold selective sodium permeability in brachiopod and annelid ASICs. All showed canonical steady-state desensitization, occurring in pH ranges higher than activating pH. Spiralian ASIC genes thus encode proton-gated cation channels, and ASICs are present in all major groups of bilaterians: Xenacoelomorpha, Protostomia, and Deuterostomia.

Hemichordate ASIC is expressed in peripheral cells and pharynx and mediates rapidly desensitizing excitatory currents in response to protons
Deuterostome ASIC expression and function have so far only been characterized in Olfactores (tunicates and vertebrates) [12,23,53], and relatively little is known about ASICs in the ambulacrarian lineage (echinoderms and hemichordates). Recent studies on neural development in sea urchin (Lytechinus variegatus) larvae showed that ASIC is expressed diffusely throughout the ciliary band [24]. This transcript (MH996684) is closely related to the Group B ASICs from echinoderms and hemichordates that showed proton-gated currents when expressed heterologously [12]. Here, we used the hemichordate Schizocardium californicum to characterize the combined expression and function of an ambulacrarian ASIC. S. californicum ASIC expression was visible in late larval stages when structures such as the nervous system and the ciliary bands become more intricate, like the sea urchin ASIC. The tornaria larva of S. californicum has two main ciliary bands: the circumoral band for feeding; and the telotroch, innervated by serotonergic neurites, for locomotion [54]. S. californicum ASIC expression was detected in both ciliary bands (Fig 5A, black arrowheads) as well as in the pharynx ( Fig  5A, orange arrowhead), however, signal intensity was variable between specimens and sometimes difficult to detect in all domains. When heterologously expressed, S. californicum ASIC showed rapidly activating and desensitizing currents in response to extracellular acidification with proton sensitivity comparable to other hemichordate ASICs but lower than echinoderm ASICs (Fig 5B, pH 50 = 5.3 ± 0.1) [12]. The S. californicum channel showed relatively weak ion selectivity, with P Na+ /P K+ values slightly above unity (2.9 ± 0.5; Fig 5D), similar to ion selectivity in previously described ambulacrarian ASICs [12]. S. californicum thus presents a functional ASIC that is expressed in the peripheral nervous system and the digestive system, and it appears that such peripheral expression of ASICs is a conserved feature of Deuterostomia, Protostomia, and Xenacoelomorpha. Filled symbols: mean (± SEM) normalized current amplitude in response to increasing proton concentrations (activation, "act.", n = 5). Open symbols: mean (± SEM) normalized current amplitude in response to pH 4 following pre-incubation in decreasing pH (steady state desensitization, n = 4).
Finally, we questioned if ASICs from Group A and Group B clades show characteristic functional differences. In terms of ion selectivity, Group B ASICs in deuterostomes show low sodium/potassium selectivity (P Na+ /P K+ between one and three), whereas Group B ASICs in protostomes and numerous Group A ASICs from deuterostomes and xenacoelomorphs have ~10-fold preference for sodium ions. In terms of apparent affinity for protons, large variability within Groups A and B is observed, with pH 50 values for activation ranging from ~4 to ~8 in both groups. Thus, we see no convincing evidence of functional characteristics that distinguish Group A and Group B ASICs.

The emergence of ASICs
Our results show that ASICs are present in Xenacoelomorpha and Protostomia in addition to Deuterostomia and are thus conserved in the three major groups of Bilateria. We also find, consistent with previous studies, that ASICs are absent from the other major lineages of Cnidaria, Placozoa, Porifera, and Ctenophora, despite the presence of other DEG/ENaC genes. This indicates that bona fide ASICs emerged in the lineage to the Bilateria, soon after the Cnidaria/Bilateria split ~800 Mya [55-57]. The closest relatives of ASICs within the DEG/ENaC tree include bile acid-gated channels (BASICs), certain peptide-gated channels (HyNaC), and a recently described leak channel (TadNaC6). BASICs are highly expressed in mammal intestines [58], HyNaCs are mostly expressed in the base of the tentacles near the mouth of the cnidarian Hydra vulgaris [11]; and TadNaC6 is expressed throughout the placozoan Trichoplax adhaerans [25]. Although none of these channels are activated by protons, the respective activity of BASICs and TadNaC6 is inhibited by increased proton concentrations ( [25,59]; S1 Fig). Although statistical support for the distinct ASIC clade is strong, we can unfortunately not resolve the precise relationships between HyNaC, BASIC, and ASIC clades, as branch support toward the base of the HyNaC+BASIC+ASIC group is relatively low (S1 Fig). Nonetheless, we think the most likely scenario is that the first ASIC emerged from a gene encoding another ligand-gated channel that was inhibited by protons. Golubovic et al. [60] have suggested that most DEG/ENaC channels, including ASICs, derive from a peptide-gated channel, noting that mammalian ASIC function is also modulated by cnidarian neuropeptides.
We infer that early bilaterians had two ASIC genes of similar proton-gated excitatory channel function, corresponding to Group A and Group B ASICs. Subsequently, and at different times, most descendants lost one of these (Fig 6). Acoels lost Group B ASIC and xenoturbellans lost Group A ASIC soon after this early split within the Xenacoelomorpha. In contrast, the first deuterostomes and, even more recently, the first chordates likely retained both ASICs, reflected in the continued presence of both in Cephalochordata (Fig 2). Subsequently, in Olfactores, Group B ASIC was lost and Group A ASIC underwent independent radiations, leading to e.g. ASIC1-ASIC4 paralogues in vertebrates [61], two Group A ASICs in tunicates [12], and multiple Group A ASICs in Cephalochordates (Fig 2).

Peripheral and central roles for ASICs
Although a broad division of the nervous system in numerous animals into central and peripheral is perhaps oversimplified [62], our results point toward ASIC expression in two distinct domains: the central nervous system, including brain and/or nerve cords; and in peripheral cells, in some cases epithelia and in other cases peripheral neurons. This points to at least two functions of ASICs: a central signaling role, where ASICs mediate rapid excitatory signals between neurons in the brain [21]; and a peripheral sensory/modulatory role, where ASICs convert increased proton concentrations in the environment-extra-organismal or in the gut-into excitatory neuronal signals [63]. In Xenacoelomopha, we found central ASICs are suitably expressed to mediate signals between nerve cord neurons and the brain or between neurons within the brain, supported by high expression of synaptotagmin in the I. pulchra brain lobes and proximal nerve cords [33], where ASIC is expressed. This central role has been best characterized in the mouse brain, where presynaptic release of acidic glutamatergic vesicles leads to rapid increases in the concentration of both glutamate and protons in the synapse, activating postsynaptic glutamate receptors and ASICs to depolarize postsynaptic neurons [21,22,64]. High expression of zebrafish ASICs in the CNS suggests a similar role in synaptic communication in other vertebrates [61]. We observed central ASICs in only one of the three Spiralia tested, and these showed projections towards the very anterior of the animal, perhaps indicative of a less integrative and more sensory role. Thus, brain ASICs are common to Xenacoelomorpha and Deuterostomia and are less prevalent in Protostomia, and this central role is more commonly played by Group A ASICs than Group B ASICs (Fig 6).
The second and peripheral domain, observed in five of the six species of Xenacoelomorpha and Spiralia studied here, largely corresponds to the ciliation in the species. While xenacoelomorphs are completely covered with ciliated epithelial cells, in many other Bilateria the ciliation is restricted to ciliary bands or the ciliary ventral region, both used for locomotion. These ASICs are thus expressed in a location to sense external changes in pH and directly modulate ciliary movement via excitatory current or, via depolarization and synaptic release onto more central neurons, send the information centrally. Regarding the latter possibility, activation of ASICs by protons can generate action potentials in cultured mouse spinal neurons [40], but whether ASIC activation could trigger synaptic release from such sensory cells, or whether xenacoelomorph ciliary cells are capable of pre-synaptic signaling like chordate sensory cells [65], is unknown. Conversely, if these ASICs are expressed on the proximal side of the epithelial cells, their role may be modulation of ciliary function in response to efferent modulatory signals. Although activation of postsynaptic ASICs has so far been linked to only glutamatergic and GABAergic synapses [21,66] and innervation of ciliary cells appears serotonergic in Spiralia [44], monoaminergic vesicles are also acidic and could thus foreseeably release protons to activate ASICs here [67].
We were surprised to see expression of ASIC predominantly in the digestive system of the spiralians O. fusiformis and P. harmeri. In mammals, chemosensors such as ASIC and transient receptor potential channels (e.g. TRPV1) are expressed in sensory neurons innervating the gastrointestinal tract [17], but oesophageal, intestinal, and lung epithelial cells also express ASICs, potentially contributing to acidinduced secretions, transport, and inflammation [68][69][70]. The gastrointestinal expression of ASICs may reflect a chemosensory role, like that mediated by ASICs in neurons innervating the lungs and skin of mammals [71,72]. Similar functions are suggested by ASIC expression in the peripheral nervous system of other chordates and Ambulacraria ( [24,73], Fig 5). Again, we can't exclude the possibility that these peripheral ASICs are modulating ciliary function, either directly via acid-induced activation or indirectly, via efferent signals from central neurons. Nonetheless, these expression patterns seem to suggest that ASICs play a sensory role in the peripheral nervous system of Deuterostomia, Protostomia, and Xenacoelomorpha via expression in (basi-)epithelial or peripheral nervous system neurons, indicating that such a role was probably present in the ancestral bilaterian. Consistent with the early appearance of this role, it is played by both Group A and Group B ASICs, which both emerged in early Bilateria. Subsequently, ASICs -particularly Group A -were likely deployed to the other, more central role for inter-neuronal communication in lineages such as Acoela and Chordata.

Loss of ASICs
The conservation and high expression of ASICs in bilaterians begs the questions as to why they were lost in certain lineages. The reason for the loss of ASICs in selected Spiralia, such as molluscs and platyhelminthes is unclear, especially when one considers that numerous molluscs are susceptible to the tide and thus vast fluctuations in pH. Presumably, gene radiations that increased sensory ion channel or transporter diversity in such animals have compensated for the loss of ASICs [74][75][76][77][78], and indeed, certain members of other channel families are capable of mediating excitation (or inhibition) in response to decreased pH [3]. Perhaps most noticeable, however, is the absence of ASICs in Ecdysozoa, a broad lineage including pan-arthropods, nematodes, and priapulids, each of which we considered in our phylogenetic analysis. Ecdysozoa split from Spiralia 600-800 Mya [55-57], adopting a chitinous cuticle utilizing rigid locomotory structures and requiring periodic molting for growth [79,80]. Concomitantly, Ecdysozoa lost the ectodermal, motile ciliated cells inherited from the last common ancestor of Cnidaria and Bilateria that mediate locomotory, feeding, secretory, and sensory functions in most other bilaterians [81,82]. The correlation between loss of ASICs and loss of motile ciliated cells, together with the conservation of other ciliated and sensory cells in Ecdysozoa, indicates that the role of ASICs in early Protostomia was primarily associated with ectodermal ciliated cells rather than more central sensory cells associated with e.g. mouth, eyes, and antennae. This, together with the presence of ASICs in the periphery of Xenacoelomorpha and Deuterostomia, suggests that the role of the earliest ASIC was likely local conversion of external chemical stimuli into modulation of locomotion, conversion of central signals into modulation of locomotion, or both. Indeed, motile ciliated epithelia in human lungs are modulated by both external and central stimuli [83], although the precise contribution of ASICs in these cells is unclear [69].

Outlook
We acknowledge that future experiments might benefit from greater genomic resources from nonbilaterian animals, but our comprehensive survey of DEG/ENaC genes from Ctenophora, Porifera, Placozoa, and Cnidaria finds no ASICs in those lineages. Our study suggests that ASICs emerged in an early bilaterian, most likely in peripheral neurons or epithelia, and were gradually adopted into the central nervous system of certain complex animals. This offers a unique insight into the role of LGICs in early bilaterian evolution. Functional characterization of diverse ASICs shows a considerable breadth of pH sensitivity and ion selectivity throughout the family, offering new tools for probing the biophysical mechanisms of function. The combined use of gene expression and function is thus a useful tool in understanding protein evolution and function.  [85], variable N-and C-termini were removed and highly similar sequences were not considered (S1 Text), and homologies were assigned by phylogenetic tree analyses based on Maximum Likelihood (ML) inferences calculated with PhyML v3.0 [86]. Robustness of tree topologies was assessed under automatic model selection based on Akaike Information Criteria. Due to computational load limitation of bootstrap performance, branch support was inferred by fast likelihoodbased methods aLRT SH-like and aBayes [87].

Animal collections and fixations
Stable cultures of acoels were maintained in the laboratory. Convolutriloba macropyga [88] were reared in a tropical aquarium system with salinity 34+/-1 ppt at a constant temperature of 25 °C. The aquariums were illuminated (Pacific LED lamp WT470C LED64S/840 PSU WB L1600, Philips) on a day/night cycle of 12/12 hours. The worms were fed with freshly hatched brine shrimp Artemia twice per week. Isodiametra pulchra [89] were cultured as described by [90], and Hofstenia miamia [91] as described by [92]. For the remaining species, adult gravid animals were collected from Bodega Bay, California, USA (Phoronopsis harmeri [93]), San Juan Island, Washington, USA (Terebratalia transversa [94]), Station Biologique de Roscoff, France (Owenia fusiformis [95]), and Kanangra Boyd National Park and Morro Bay State Park, California, USA (Schizocardium californicum [96]). Animals were spawned and larvae obtained as described in [97][98][99][100]. Adult and larval specimens were starved for 2 -7 days prior to fixation. Samples were relaxed in 7.4% magnesium chloride and fixed in 4% paraformaldehyde in culture medium for 1 hour at room temperature and washed several times in 0.1% Tween 20 phosphate buffered saline (PBS), dehydrated through a graded series of methanol, and stored in pure methanol or ethanol at −20 °C.

Cloning
The full-length coding sequences of identified ASIC genes were amplified from cDNA of C. macropyga, I. pulchra, H. miamia, P. harmeri, O. fusiformis, T. transversa, and S. californicum by PCR using gene-specific primers. PCR products were purified and cloned into a pGEM-T Easy vector (Promega, A1360) according to the manufacturer's instructions and the identity of inserts confirmed by sequencing. Riboprobes were synthesized with Ambion Megascript T7 (AM1334) and SP6 (AM1330) kit following manufacturer's instruction for subsequent in situ hybridization.

Immunohistochemistry and situ hybridization
Single whole-mount colorimetric and fluorescent in situ hybridization was performed following an established protocol [31] with probe concentration of 0.1 ng/μl (I. pulchra) or 1 ng/μl (the remaining species) and hybridization temperature of 67 °C. Proteinase K treatment time was adjusted for each species and ranged from 2 minutes (P. harmeri, O. fusiformis, S. californicum) to 10 minutes (T. transversa). Post-hybridization low salt washes were performed with 0.05x saline sodium citrate (SSC; H. miamia) or 0.2x SSC (the remaining species). Fluorescent in situ hybridization was visualized with TSA Cy3 kit (PerkinElmer, NEL752001KT). Samples were mounted in 70% glycerol or subjected to immunohistochemistry for visualization of neural structures: samples were permeabilized in 0,2 % Triton-x in PBS (PTx) and blocked in 1% bovine serum albumin in PTx (PBT) and incubated with antibodies against tyrosinated tubulin (Sigma, T9028) at a concentration of 1:250 in PTx with 5% normal goat serum, and incubated for 16-18 hours at 4 °C. After several washes in PBT the samples were incubated with secondary goat anti-mouse antibodies conjugated with AlexaFluor 488 (Life Technologies), at a concentration 1:200 in PTx with 5% normal goat serum for 16-18 hours at 4 °C, and samples washed extensively before mounting in 70 % glycerol and imaging. Nuclei were stained with DAPI (Molecular Probes).

Imaging
Representative specimens from colorimetric in situ hybridization experiments were imaged with a Zeiss Axiocam 503 color connected to a Zeiss Axioscope 5 using bright-field Nomarski optics. Fluorescently labelled samples were scanned in an Olympus FV3000 confocal laser-scanning microscope. Colorimetric in situ's stained with antibodies were scanned in a Leica SP5 confocal laser-scanning microscope using reflection microscopy protocol as described by [101]. Images were analyzed with Imaris 9.8.0 and Photoshop CS6 (Adobe), and figure plates were assembled with Illustrator CC. Brightness/contrast and color balance adjustments were applied to the whole image, not parts.

Electrophysiological recordings and data analysis
For expression in Xenopus laevis oocytes and electrophysiological experiments, coding sequences were mutated synonymously to remove internal restriction sites if necessary and subcloned into SalI and XbaI sites of a modified pSP64poly(A) vector (ProMega), containing 5' SP6 sequence, 5′-and 3′-UTR sequences of the X. laevis β-globin gene, and a C-terminal Myc tag, with an EcoRI restriction site after the poly(A) tail (S2 Text). Lingula anatina ASIC (g20471.t1 from Lingula anatina Ver 2.0, OIST Marine Genomics Unit), synonymously mutated to remove internal restriction sites and including a Cterminal myc tag before the stop codon, was commercially synthesized and subcloned (Genscript) into HindIII and BamHI sites of pSP64poly(A) (S2 Text). Plasmids were linearized with EcoRI, and cRNA was synthesized in vitro with SP6 Polymerase (mMessage mMachine kit, Ambion). Stage V/VI X. laevis oocytes, purchased from Ecocyte Bioscience (Dortmund, Germany), were injected with 3-90 ng cRNA. After injection, oocytes were incubated for one to three days at 19°C in 50% Leibowitz medium (Merck) supplemented with 0.25 mg/ml gentamicin, 1 mM L-glutamine, and 15 mM HEPES (pH 7.6). Whole cell currents were recorded from oocytes by two-electrode voltage clamp using an OC-725C amplifier (Warner Instruments) and an LIH 8+8 digitizer with Patchmaster software (HEKA), acquired at 1 kHz and filtered at 200 Hz. Currents were also analyzed in pClamp v10.7 software (Molecular Devices) and additionally filtered at 1 Hz for display in figures. Oocytes were clamped at -60 mV, unless otherwise indicated, and continuously perfused with a bath solution containing (in mM): 96 NaCl, 2 KCl, 1.8 CaCl 2 , 1 MgCl 2 , and 5 HEPES (for pH > 6.0) or 5 MES (for pH ≤ 6.0). pH was adjusted with NaOH, HCl, or KOH, as appropriate. In most experiments, activating/desensitizing pH was applied to oocytes in between resting periods (at pH 7.5 for most ASICS, at pH 9.0, 8.6, and 8.0 for O. fusiformis, L. anatina S. californicum ASICs, respectively, unless otherwise indicated) of at least 30 s. After retrieving current amplitude from pClamp, all data analyses were performed in Prism v9 (GraphPad Software). In concentration-response graphs, currents are normalized to maximum protongated current. For ion selectivity experiments, IV relationships were measured in regular bath solution and that in which extracellular NaCl was replaced with KCl. IV relationships were obtained by activating the channels at different membrane potentials from -80 to 60 mV, with 20 mV increments, unless otherwise indicated. Reversal potentials (V rev,Na+ and V rev,K+ ) were taken from the intersection of the IV curve with the voltage axis. These values were used to calculate relative permeability P Na+ /P K+ with the Goldman-Hodgkin-Katz equation, P Na+ /P K+ = exp(F(V rev,Na+ -V rev,K+ )/RT), where F = Faraday constant, R = gas constant, and T = 293 K. Standard chemicals were purchased from Merck. Specialist chemicals (S1 Fig) were purchased from Santa Cruz Biotechnology (item sc-222407, sodium ursodeoxycholic acid, ≥98% purity) or synthesized by Genscript ((pyroE)WLGGRFamide, ≥97% purity -"Hydra RFamide I" from [11]).