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phenotype, or whether it acts in a dominant or recessive fashion. The recent systematic analysis of mouse knockouts (complete abrogation of gene function) reports that 35% (56 of 160) appeared completely normal [18]. In other words, even with the application of rigorous criteria to define a highly deleterious mutation, we can still find these supposedly pathogenic mutations in healthy people. The reluctance of schizophrenia researchers to claim they have found causal mutations is understandable. Just what will it take to get to the point of declaring victory? The scale of the problem is made clear in a recent discovery of protein-altering variants in SLC30A8, confirming that the gene is involved in type 2 diabetes. Analysis of about 150,000 individuals was needed to secure the finding [19]. We are nowhere near that number for schizophrenia; indeed, it’s possible we’ll never have the type of evidence that we have for genes involved in diabetes or inflammatory bowel disease. The flexibility and adaptability that is such a notable feature of human behaviour brings a freedom from genetic determination that presumably extends to behaviour in illness. While our understanding of the genetic architecture of behaviour is too limited to make definitive claims, there is a real possibility that the sought after large effect mutations may be of much smaller effect than is hoped for. New approaches may be needed to work up the biological implications of our hard won insights into the genetic basis of psychiatric disease.

References 1. Ustun, T.B., Ayuso-Mateos, J.L., Chatterji, S., Mathers, C., and Murray, C.J. (2004). Global burden of depressive disorders in the year 2000. Br. J. Psychiatry 184, 386–392. 2. Schizophrenia Working Group of the Psychiatric Genomics Consortium (2014). Biological insights from 108 schizophrenia associated genetic loci. Nature 511, 421–427. 3. Flint, J., and Kendler, K.S. (2014). The genetics of major depression. Neuron 81, 484–503. 4. Purcell, S.M., Wray, N.R., Stone, J.L., Visscher, P.M., O’Donovan, M.C., Sullivan, P.F., and Sklar, P. (2009). Common polygenic variation contributes to risk of schizophrenia and bipolar disorder. Nature 460, 748–752. 5. Wellcome Trust Case Control Consortium (2007). Genome-wide association study of 14,000 cases of seven common diseases and 3,000 shared controls. Nature 447, 661–678. 6. Newton-Cheh, C., Johnson, T., Gateva, V., Tobin, M.D., Bochud, M., Coin, L., Najjar, S.S., Zhao, J.H., Heath, S.C., Eyheramendy, S., et al. (2009). Genome-wide association study identifies eight loci associated with blood pressure. Nat. Genet. 41, 666–676. 7. Fromer, M., Pocklington, A.J., Kavanagh, D.H., Williams, H.J., Dwyer, S., Gormley, P., Georgieva, L., Rees, E., Palta, P., Ruderfer, D.M., et al. (2014). De novo mutations in schizophrenia implicate synaptic networks. Nature 506, 179–184. 8. Purcell, S.M., Moran, J.L., Fromer, M., Ruderfer, D., Solovieff, N., Roussos, P., O’Dushlaine, C., Chambert, K., Bergen, S.E., Kahler, A., et al. (2014). A polygenic burden of rare disruptive mutations in schizophrenia. Nature 506, 185–190. 9. Nejentsev, S., Walker, N., Riches, D., Egholm, M., and Todd, J.A. (2009). Rare variants of IFIH1, a gene implicated in antiviral responses, protect against type 1 diabetes. Science 324, 387–389. 10. Rivas, M.A., Beaudoin, M., Gardet, A., Stevens, C., Sharma, Y., Zhang, C.K., Boucher, G., Ripke, S., Ellinghaus, D., Burtt, N., et al. (2011). Deep resequencing of GWAS loci identifies independent rare variants associated with inflammatory bowel disease. Nat. Genet. 43, 1066–1073. 11. Girard, S.L., Gauthier, J., Noreau, A., Xiong, L., Zhou, S., Jouan, L., Dionne-Laporte, A., Spiegelman, D., Henrion, E., Diallo, O., et al. (2011). Increased exonic de novo mutation rate in individuals with schizophrenia. Nat. Genet. 43, 860–863. 12. Xu, B., Roos, J.L., Dexheimer, P., Boone, B., Plummer, B., Levy, S., Gogos, J.A., and

Evolution: Ctenophore Genomes and the Origin of Neurons Recent sequencing of ctenophore genomes opens a new era in the study of this unique and phylogenetically distant group. The presence of neurodevelopmental genes, pre- and postsynaptic modules, and transmitter molecules is consistent with a single origin of neurons. Heather Marlow and Detlev Arendt The origin of neurons and nervous systems is one of the most exciting questions in animal evolution. Neurons, electrically excitable cells that signal to target cells via synapses, are

found in three animal lineages: in the bilaterians — comprising vertebrates, insects, nematodes and other groups often found with ganglia, nerve cords and brains; in the cnidarians — polyps and jellyfish with nerve nets that cover the entire body; and in a third








Karayiorgou, M. (2011). Exome sequencing supports a de novo mutational paradigm for schizophrenia. Nat. Genet. 43, 864–868. Xu, B., Ionita-Laza, I., Roos, J.L., Boone, B., Woodrick, S., Sun, Y., Levy, S., Gogos, J.A., and Karayiorgou, M. (2012). De novo gene mutations highlight patterns of genetic and neural complexity in schizophrenia. Nat. Genet. 44, 1365–1369. Nusslein-Volhard, C., and Wieschaus, E. (1980). Mutations affecting segment number and polarity in Drosophila. Nature 287, 795–801. Metzstein, M.M., Stanfield, G.M., and Horvitz, H.R. (1998). Genetics of programmed cell death in C. elegans: past, present and future. Trends Genet. 14, 410–416. MacArthur, D.G., Manolio, T.A., Dimmock, D.P., Rehm, H.L., Shendure, J., Abecasis, G.R., Adams, D.R., Altman, R.B., Antonarakis, S.E., Ashley, E.A., et al. (2014). Guidelines for investigating causality of sequence variants in human disease. Nature 508, 469–476. MacArthur, D.G., Balasubramanian, S., Frankish, A., Huang, N., Morris, J., Walter, K., Jostins, L., Habegger, L., Pickrell, J.K., Montgomery, S.B., et al. (2012). A systematic survey of loss-of-function variants in human protein-coding genes. Science 335, 823–828. White, J.K., Gerdin, A.K., Karp, N.A., Ryder, E., Buljan, M., Bussell, J.N., Salisbury, J., Clare, S., Ingham, N.J., Podrini, C., et al. (2013). Genomewide generation and systematic phenotyping of knockout mice reveals new roles for many genes. Cell 154, 452–464. Flannick, J., Thorleifsson, G., Beer, N.L., Jacobs, S.B., Grarup, N., Burtt, N.P., Mahajan, A., Fuchsberger, C., Atzmon, G., Benediktsson, R., et al. (2014). Loss-of-function mutations in SLC30A8 protect against type 2 diabetes. Nat. Genet. 46, 357–363.


Trust Centre for Human Genetics, University of Oxford, Oxford OX3 7BN, UK. 2MRC Integrative Epidemiology Unit, UK Centre for Tobacco and Alcohol Studies, and School of Experimental Psychology, University of Bristol, BS8 1TU, UK. *E-mail: [email protected]


group that, until very recently, has received little attention — the enigmatic ctenophores, or ‘comb jellies’ (Figure 1A). The ctenophore nervous system is a nerve net (Figure 1B) with local aggregations of neurons, most pronounced around the apex of the animal [1,2] (Figure 1C). As the gelatinous body of the ctenophores resembles that of cnidarian jellyfish, many authors assumed that cnidarians and ctenophores are related (grouped as ‘coelenterates’ [3]). Yet, this view has been challenged by the very different way these animals move: while rhythmic muscle contractions propel the cnidarian medusae, the comb jellies

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Figure 1. The ctenophore body plan and nervous system. (A) General organization of a comb jelly. Drawing of Pleurobrachia pileus taken from [9]. (B) Ctenophore nerve net. (Reproduced with permission from Macmillan Publishers Ltd [1]). (C) Nervous system around the aboral sensory organ, reproduced from [2]. AO, apical organ; Cg, ciliated groove; Pf, polar field. Adults of Pleurobrachia bachei stained for an antibody directed against tyrosinated alpha-tubulin.

swim by means of myriads of cilia, arranged in characteristic rows of comb plates (Figure 1A). Others considered the ctenophores to be more closely related to the bilaterians, as they have mesoderm and true muscle cells. Despite disagreement on the precise branching order, however, it has so far been generally accepted that bilaterians, cnidarians and ctenophores should be more closely related to each other than to sponges and placozoans, two groups of animals with very simple, amorphous morphology and few cell types, lacking bona fide neurons and muscle cells [4]. Sponges and placozoans are commonly regarded to have been the first of these ‘basal metazoan’ groups to have branched off the animal tree of life. To infer the step-wise assembly of the nervous system in animal evolution [5], we need first to understand the branching order of the ‘basal metazoan’ lineages; and second, a detailed account of neural

characteristics present in each lineage, at the level of genes, cell types, tissue and, ultimately, behavior. Recent sequencing of basal metazoan genomes [1,6,7] represents a significant advance in both areas, providing new insight into the early branching of the metazoan phylogenetic tree and allowing us to reconstruct the emergence of the components of key cellular modules characteristic of neurons and nervous systems. Analyses of the recently published genomes of the comb jellies Mnemiopsis leydii [7] and Pleurobrachia [1] (Figure 1A) now come as a particular surprise. Aligning the ctenophore predicted protein sequences to those of other animal groups and calculating the phylogenetic trees most consistent with the observed sequence divergence, both studies seem to find that the ctenophore lineage (and not sponges) represents the earliest branch of the animal tree of life. The authors argue by extension that the diverse array of ctenophore cell types (including neurons and muscle) must have arisen independently from those found in later branching animal lineages (cnidarians and bilaterians), resulting in ‘‘extensive parallel evolution of neural organization’’ [1]. Moroz et al. [1] set out to test this scenario through presence/absence analysis of gene families and interpret the results of their analysis as supporting independent evolution of neurons. They also conduct a limited experimental characterization of the ctenophore nervous system and find that glutamate plays a major role as a neurotransmitter, triggering the contraction of muscle cells; they find that other transmitters do not seem to be involved. The authors interpret this as an ancient condition, dissimilar from that of other nervous systems. While a finding supporting the independent evolution of nervous systems would indeed be remarkable, we have to step back and assess the implications of the new ctenophore data separately with regard to the two points mentioned above. What do we learn about the branching order of basal metazoans? And what does the complement of ‘neural genes’ present in ctenophores reveal about nervous system origins? Regarding the phylogenetic placement of ctenophores, a note of caution seems to be warranted. A previous molecular phylogenetic

study [3] using refined evolutionary models for sequence divergence had suggested that long-branch attraction might be responsible for the basal position of the ctenophores. Long-branch attraction is an artefact in which species which have undergone a rapid and extensive amount of change from the ancestral state — resulting in a ‘long branch’ in the phylogenetic tree — are attracted to other ‘long branches’ or, in this case, the long root of the animal tree [3]. Worryingly, the inclusion of additional ctenophore transcriptomes, which should allow clearer placement of the group, abolishes support for the basal placement [1], so that the available evidence appears to be insufficient to ultimately and convincingly place the ctenophores. (In Figure 2A, the ctenophore branch is tentatively placed between that of placozoans and cnidarians — representing only one of several solutions consistent with the available phylogenetic data, while other possible solutions are indicated by dashed lines.) If the phylogenetic signal remains inconclusive, what can we learn about nervous system origins from the presence or absence of specific genes? ’Neural genes’ are of two kinds: developmental genes involved in nervous system specification or morphogenesis, and differentiation genes encoding neuron-specific structures and functions. Among the developmental genes, Moroz et al. [1] specifically cite the absence of neuroD and neurogenin transcription factors. However, these genes are uninformative as they evolved only in bilaterians [8]. Importantly, they do not discuss the SoxB [9] and LIM homeodomain genes [10], which are well-known for their role in neuron type specification and, consistent with this, are prominently expressed in ctenophore neural tissue such as the apical sensory organ [9,10], strongly indicative of conserved roles in nervous system development. Thus, in contrast to what Moroz et al. [1] state, the limited knowledge available on ctenophore neurodevelopmental genes appears largely consistent with nervous system homology — but, admittedly, does not prove it, as the same transcription factors also exist in the neuron-less sponges. Another test for nervous system homology is to survey the presence of genes that encode structures and

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functions that are characteristic and, in the ideal case, specific for neurons. Recent comparative genomic and transcriptomic studies in basal metazoans, including ctenophores, have thus focused on the presence or absence of pre- and postsynaptic genes [1,6,7]. Within the synapse, the postsynapse receives the transmitter signal sent by the presynapse. Postsynaptic genes are present in ctenophores [1], but the sequencing of a sponge genome [6] had also identified an almost complete gene complement of the ‘postsynaptic density’, indicating that the emergence of this cellular module predated the evolution of neurons with morphological synapses [5] and is thus uninformative with regard to nervous system homology. One hypothesis explaining this counterintuitive finding is that the ‘postsynapse’ is a modified sensory receptive field that acquired the capacity to detect intercellular signals [5,11]. This would also account for the recruitment of glutamate as a transmitter (extensively used in ctenophores [1]): glutamate might have initially been detected as an environmental signal, later facilitating its recruitment as intercellular transmitter following the advent of the synapse [5,11]. In sponges, postsynaptic density proteins are indeed localized to flask-shaped sensory cells [12]; furthermore, as glutamate has been reported to trigger contractions as an intercellular paracrine signal in sponges [13], ‘postsynaptic’ receptor assemblies may already be involved in the propagation of contraction waves. If the postsynapse is not specific for neurons, what can be said about the presynapse? The emergence of the presynapse allowed direct and targeted information transfer to the postsynapse of other cells and may indeed represent the key novelty in neuron evolution. Ultrastructurally, the ctenophore presynapse exhibits a unique morphology, referred to as the presynaptic triad (Figure 2B), stereotypically composed of a mitochondrion, an extension of the endoplasmic reticulum and synaptic vesicles facing the membrane [14]. This peculiar morphology, however, does not preclude presynapse homology, given that the same cellular components are also involved in presynaptic morphology in bilaterians (Figure 2C) [15]. Transmitter release

x SNAP-25


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Figure 2. The phylogenetic position of ctenophores and the evolution of neurons. (A) The step-wise assembly of synapses in metazoan evolution. Dashed lines demarcate possible positions of the ctenophore branch in a simplified animal evolutionary tree. Boxes indicate the gain of a cellular module or its constituting proteins. Coloured bars demarcate animal groups that possess the proteins labelled with the same colour code in panel D. Animal drawings from [4]. (B) Diagram of the ctenophore presynaptic triad. sER: smooth endoplasmic reticulum; v: vesicle; m: mitochondrium. From [14] with kind permission from Springer Science and Business Media. (C) Diagram of the vertebrate presynapse. sER, smooth endoplasmic reticulum; v, vesicle; m, mitochondrium. From [15]. (D) The presynaptic active zone protein complex. Colored proteins form the evolutionarily conserved core of active zones [16]. Colour code refers to the phylogenetic occurrence of individual proteins as indicated by bars in panel A. Adapted from [16]. (E) Fiber cells in Trichoplax adherens after https://sites.google.com/a/ poriferaproject.com/www/moretrichoplaxadhaerens; s: synpase-like intercellular connection with vesicles (v); b: intracellular bacterium; k: concrement ‘vacuole’; m: mitochondria.

occurs from the presynaptic ‘active zone’, which aligns with the postsynaptic density [16]. Among the

active zone proteins (Figure 2D) some have more general functions outside the synaptic context and are thus

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uninformative to track synapse origins; for example, the SNARE complex components (including synaptobrevin, and syntaxin) generally mediate vesicle fusion and also exist in unicellular eukaryotes [6,16]. Not surprisingly, these proteins are present in the ctenophore genomes [1]. Similarly, synaptotagmin, which is found in all animals, including those without nervous systems, functions more generally in calcium-mediated vesicle exocytosis. Three of the more general CAZ proteins appear to be absent in Pleurobrachia, such as profilin, an actin-binding protein generally involved in restructuring of the actin cytoskeleton or synaptogyrin, a membrane protein involved in modulating vesicle exocytosis [1]. Both of these genes have, however, been identified and annotated in Mnemiopsis [7] and thus represent secondary losses restricted to a subset of the ctenophore lineage. Key to tracking presynapse origins are five proteins that form the core of active zones (bold black outlines in Figure 2D) [16]. RIM multidomain proteins are central organizers of active zones; for example, the Rab3-RIM-Munc13 complex brings synaptic vesicles in close proximity to the priming machinery [16]. ELKS is another important matrix protein with multiple binding partners required for the correct localization of other active zone components [16]. These proteins (not discussed by Moroz et al. [1]) thus represent excellent candidates for tracking the evolution of the presynapse; and consistent with the absence of synapses in sponges, RIMs and ELKS are reported to be absent from the sponge genomes [6]. If these proteins are present in the ctenophore presynapse, homology of synapses — and thus of neurons and nervous systems between ctenophores and other animals — will be strongly supported. Another argument that Moroz et al. [1] put forward in support of the unique nature of the ctenophore nervous system is the apparent paucity of transmitter molecules. Recording from single muscle cells, they make a strong case that glutamatergic transmission alone governs the contraction of muscles [1] (consistent with glutamate being involved in the propagation of contractile waves in sponges [13] and playing a predominant role in muscular control in other animals). However,

these recordings fail to uncover the transmitter utilized in the nerve nets [2] and the ciliomotor systems [17]. Previous pharmacological and electrophysiological studies had made a strong case for cholinergic transmission in the control of ciliary beating and luminescence in ctenophores [18], conflicting with the new finding that muscarinic and nicotinic acetylcholine receptors appear to be missing from Pleurobrachia [1]. Besides glutamate and acetycholine, GABA may also play a role [1], as supported by the presence of a metabotropic GABAB receptor in Mnemiopsis [7] and consistent with the effect of GABA on sponge contractile behavior [13]. Thus, the set of transmitters employed by the ctenophore nervous system might be more extended than Moroz et al. [1] suggest. Finally, Moroz and colleagues [1] focus on the neuron-specific, Elav-like RNA-binding proteins that regulate RNA splicing and abundance in neurons to control transmitter levels and neuronal excitability [1]. They report expression of one of three paralogs in cells in the comb plate, where the multiple ciliated cells reside that drive the characteristic ciliary swimming. The authors claim that ‘‘in Pleurobrachia Elav has not been detected in neural tissues’’ [1]; however, given that the electrically coupled ciliomotor cells of the comb plates respond to synaptic input from the ectodermal nerve net via action potentials [17], it is not unexpected to find an Elav-like gene expressed in these cells. In any case, it will be interesting to identifiy the expression of the other Elav paralogs in ctenophores. The new ctenophore genomes [1,7] are significant contributions to our toolkit for early metazoan evolutionary studies and, in particular, for understanding nervous system origins. If ctenophores are indeed positioned basally to cnidarians, as suggested by these studies, they represent an important link from pre-neural metazoan forms to the neuron-bearing cnidarians and bilaterians. If, ultimately, ctenophores were found to be basal to sponges, this would be indicative of secondary simplification in the sponge lineage (a notion backed by mosaic gene loss and the presence of Hox and Parahox ghost loci [19]); in this context it will be

interesting to test for the possible mosaic presence of presynaptic components in sponge cellular transcriptomes. Also, it will be highly rewarding to test for the presence of absence of neural modules in the transcriptomes of cell types present in the placozoans [20], such as the neuron-like fibre cells with long cellular extensions and junctions that may support electrical conduction (Figure 2E). A solution to the century-old question of nervous system origins appears imminent. References 1. Moroz, L.L., Kocot, K.M., Citarella, M.R., Dosung, S., Norekian, T.P., Povolotskaya, I.S., Grigorenko, A.P., Dailey, C., Berezikov, E., Buckley, K.M., et al. (2014). The ctenophore genome and the evolutionary origins of neural systems. Nature 510, 109–114. 2. Jager, M., Chiori, R., Alie, A., Dayraud, C., Queinnec, E., and Manuel, M. (2010). New insights on ctenophore neural anatomy: immunofluorescence study in Pleurobrachia pileus (Muller, 1776). J. Exp. Zool. B Mol. Dev. Evol. 316B, 171–187. 3. Philippe, H., Derelle, R., Lopez, P., Pick, K., Borchiellini, C., Boury-Esnault, N., Vacelet, J., Renard, E., Houliston, E., and Queinnec, E. (2009). Phylogenomics revives traditional views on deep animal relationships. Curr. Biol. 19, 706–712. 4. Nielsen, C. (2012). Animal Evolution: Interrelationships of the Living Phyla, 3 Edition (Oxford: Oxford University press). 5. Achim, K., and Arendt, D. (2014). Structural evolution of cell types by step-wise assembly of cellular modules. Curr. Opin. Neurobiol. 27C, 102–108. 6. Srivastava, M., Simakov, O., Chapman, J., Fahey, B., Gauthier, M.E., Mitros, T., Richards, G.S., Conaco, C., Dacre, M., Hellsten, U., et al. (2010). The Amphimedon queenslandica genome and the evolution of animal complexity. Nature 466, 720–726. 7. Ryan, J.F., Pang, K., Schnitzler, C.E., Nguyen, A.D., Moreland, R.T., Simmons, D.K., Koch, B.J., Francis, W.R., Havlak, P., Smith, S.A., et al. (2013). The genome of the ctenophore Mnemiopsis leidyi and its implications for cell type evolution. Science 342, 1242592. 8. Simionato, E., Ledent, V., Richards, G., Thomas-Chollier, M., Kerner, P., Coornaert, D., Degnan, B.M., and Vervoort, M. (2007). Origin and diversification of the basic helix-loop-helix gene family in metazoans: insights from comparative genomics. BMC Evol. Biol. 7, 33. 9. Jager, M., Queinnec, E., Chiori, R., Le Guyader, H., and Manuel, M. (2008). Insights into the early evolution of SOX genes from expression analyses in a ctenophore. J. Exp. Zool. B. Mol. Dev. Evol. 310, 650–667. 10. Simmons, D.K., Pang, K., and Martindale, M.Q. (2012). Lim homeobox genes in the Ctenophore Mnemiopsis leidyi: the evolution of neural cell type specification. Evodevo 3, 2. 11. Shaham, S. (2010). Chemosensory organs as models of neuronal synapses. Nat. Rev. Neurosci. 11, 212–217. 12. Sakarya, O., Armstrong, K.A., Adamska, M., Adamski, M., Wang, I.F., Tidor, B., Degnan, B.M., Oakley, T.H., and Kosik, K.S. (2007). A post-synaptic scaffold at the origin of the animal kingdom. PLoS One 2, e506. 13. Ellwanger, K., Eich, A., and Nickel, M. (2007). GABA and glutamate specifically induce contractions in the sponge Tethya wilhelma. J. Comp. Physiol. A Neuroethol. Sens. Neural. Behav. Physiol. 193, 1–11.

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14. Hernandez-Nicaise, M.L. (1973). The nervous system of ctenophores. III. Ultrastructure of synapses. J. Neurocytol. 2, 249–263. 15. Droz, B., Rambourg, A., and Koenig, H.L. (1975). The smooth endoplasmic reticulum: structure and role in the renewal of axonal membrane and synaptic vesicles by fast axonal transport. Brain Res. 93, 1–13. 16. Sudhof, T.C. (2012). The presynaptic active zone. Neuron 75, 11–25. 17. Moss, A.G., and Tamm, S.L. (1987). A calcium regenerative potential controlling ciliary reversal is propagated along the length of

ctenophore comb plates. Proc. Natl. Acad. Sci. USA 84, 6476–6480. 18. Anctil, M. (1985). Cholinergic and monoaminergic mechanisms associated with the control of bioluminescence in the ctenophore Mnemiopsis leidyi. J. Exp. Biol. 119, 225–238. 19. Mendivil Ramos, O., Barker, D., and Ferrier, D.E. (2012). Ghost loci imply Hox and ParaHox existence in the last common ancestor of animals. Curr. Biol. 22, 1951–1956. 20. Smith, C.L., Varoqueaux, F., Kittelmann, M., Azzam, R.N., Cooper, B., Winters, C.A., Eitel, M., Fasshauer, D., and Reese, T.S. (2014). Novel cell types, neurosecretory cells, and

body plan of the early-diverging metazoan Trichoplax adhaerens. Curr. Biol. 24, 1565–1572.

Developmental Biology Unit, European Molecular Biology Laboratory, Meyerhofstraße 1, 69012 Heidelberg, Germany. E-mail: [email protected], [email protected] http://dx.doi.org/10.1016/j.cub.2014.06.057

Evolution: ctenophore genomes and the origin of neurons.

Recent sequencing of ctenophore genomes opens a new era in the study of this unique and phylogenetically distant group. The presence of neurodevelopme...
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