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. Author manuscript; available in PMC: 2015 Feb 23.

The Ctenophore Genome and the Evolutionary Origins of Neural Systems

Leonid L Moroz1,2,3,Kevin M Kocot4,Mathew R Citarella1,Sohn Dosung1,Tigran P Norekian1,3,Inna S Povolotskaya5,6,Anastasia P Grigorenko7,8,Christopher Dailey9,Eugene Berezikov10,Katherine M Buckley11,Andrey Ptytsyn1,Denis Reshetov8,Krishanu Mukherjee1,Tatiana P Moroz1,Yelena Bobkova1,Fahong Yu2,Vladimir V Kapitonov12,Jerzy Jurka12,Yuri Bobkov1,Joshua J Swore1,3,David O Girardo1,3,Alexander Fodor1,Fedor Gusev7,8,Rachel Sanford1,Rebecca Bruders1,3,Ellen Kittler13,Claudia E Mills3,Jonathan P Rast11,Romain Derelle5,6,Victor V Solovyev14,Fyodor A Kondrashov5,6,15,Billie J Swalla3,Jonathan V Sweedler8,Evgeny I Rogaev7,8,16,17,Kenneth M Halanych4,Andrea B Kohn1
1The Whitney Laboratory for Marine Bioscience, University of Florida, 9505 Ocean Shore Blvd., St. Augustine, Florida 32080, USA
2Department of Neuroscience & McKnight Brain Institute, University of Florida, Gainesville, FL 32611, USA
3Friday Harbor Laboratories, University of Washington, Friday Harbor, WA 98250, USA
4Department of Biological Sciences, Auburn University, 101 Rouse Life Sciences, Auburn, Alabama 36849, USA
5Centre for Genomic Regulation (CRG), Dr. Aiguader 88, 08003 Barcelona, Spain
6Universitat Pompeu Fabra (UPF), Barcelona, Spain
7Department of Psychiatry, University of Massachusetts Medical School, l303 Belmont Street, Worcester MA 01605, USA
8Vavilov Institute of General Genetics, Russian Academy of Sciences (RAS), Gubkina 3, Moscow 119991, RF
9Department of Chemistry and the Beckman Institute for Advanced Science and Technology, University of Illinois Urbana-Champaign, Urbana, IL 61801, USA
10European Research Institute for the Biology of Ageing, University of Groningen Medical Center, Antonius Deusinglaan 1, Building 3226, Room 03.34, 9713 AV Groningen; The Netherlands
11Department of Medical Biophysics and Department of Immunology, University of Toronto, Sunnybrook Research Institute 2075 Bayview Avenue, Toronto, ON M4N 3M5, Canada
12Genetic Information Research Institute, 1925 Landings Dr., Mountain View, CA 94043, USA
13Program in Molecular Medicine, University of Massachusetts Medical School, 222 Maple Avenue, Shrewsbury, Massachusetts 01545, USA
14Department of Computer Science, Royal Holloway, University of London, Egham, Surrey TW20 0EX, UK
15Institució Catalana de Recerca i Estudis Avançats (ICREA), Pg. Lluis Companys 23, 08010 Barcelona, Spain
16Center for Brain Neurobiology and Neurogenetics and Institute of Cytology and Genetics, RAS, Lavrentyev Ave., 10, Novosibirsk 630090, RF
17Faculty of Bioengineering and Bioinformatics, Lomonosov Moscow State University, Leninskiye Gory, 119991, Moscow, RF
✉

Corresponding to: Leonid L Moroz (moroz@whitney.ufl.edu), Principal Investigator; Kenneth Halanych (ken@auburn.edu), phylogenomiucs; Evgeny I. Rogaev (Evgeny.Rogaev@umassmed.edu), gDNA-seq; Andrea B. Kohn (abkohn@msn.com), RNA-seq

Issue date 2014 Jun 5.

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PMCID: PMC4337882  NIHMSID: NIHMS589169  PMID:24847885
The publisher's version of this article is available atNature

Abstract

The origins of neural systems remain unresolved. In contrast to other basal metazoans, ctenophores, or comb jellies, have both complex nervous and mesoderm-derived muscular systems. These holoplanktonic predators also have sophisticated ciliated locomotion, behaviour and distinct development. Here, we present the draft genome ofPleurobrachia bachei, Pacific sea gooseberry, together with ten other ctenophore transcriptomes and show that they are remarkably distinct from other animal genomes in their content of neurogenic, immune and developmental genes. Our integrative analyses place Ctenophora as the earliest lineage within Metazoa. This hypothesis is supported by comparative analysis of multiple gene families, including the apparent absence of HOX genes, canonical microRNA machinery, and reduced immune complement in ctenophores. Although two distinct nervous systems are well-recognized in ctenophores, many bilaterian neuron-specific genes and genes of “classical” neurotransmitter pathways either are absent or, if present, are not expressed in neurons. Our metabolomic and physiological data are consistent with the hypothesis that ctenophore neural systems, and possibly muscle specification, evolved independently from those in other animals.

Approximately 150 recognized species of comb jellies form a clade of pre-bilaterian animals1–3(Fig. 1f) with an elusive genealogy, possibly tracing their ancestry to the Ediacaran biota4,5. We selected the Pacific sea gooseberry,Pleurobrachia bachei (A. Agassiz, 1860,Fig. 1a,Extended_Data_Fig. 1,Supplementary_Data_SD1 andvideos) as a model ctenophore due to preserved traits thought ancestral for this lineage (e.g. cydippid larva and tentacles). Three next-generation sequencing platforms (454/Illumina/Ion Torrent) were used to obtain >700-fold coverage (Supplementary_Tables_1–2S) ofPleurobrachia’s genome, and about 2,000-fold coverage of the transcriptome representing all major organs and developmental stages (Supplementary_Tables_3–4S). Consequently, the draft assembly was 156,146,497 base pairs (bp) with 19,523 predicted protein-coding genes (Supplementary_Tables_5–7S). About 90% of these predicted genes are expressed in at least one tissue or developmental stage (Supplementary_Methods) and 44% ofPleurobrachia genes have orthologs in other animals (Supplementary_Tables_7–8S). More than 300 families of transposable elements (TEs) constitute at least 8.5% of the genome (Supplementary_Table_9S,Supplementary_Data_SD2) with numerous examples of diversification of some ancient TE classes (e.g. transposases, reverse transcriptases, etc). Approximately 1.0% of the genome is methylated.Pleurobrachia also employs DNA demethylation during development, with both 5-hydroxymethyl cytosine (5hmC) and its synthetic enzyme TET6 (Extended_Data_Fig. 2). The obtained genome and transcriptome data provide rich resources (http://moroz.hpc.ufl.edu/) for investigating both animal phylogeny and evolution of animal innovations including nervous systems2,3,7–9.

Figure 1. Ctenophores and their innovations.

Figure 1

a, The sea gooseberry,Pleurobrachia bachei (Fig. 1S) was selected as a target for genome sequencing due to preservation of traits ancestral for this lineage and sincein situ hybridization/immunolabeling is possible. b–d: Major ctenophore innovations.b, Nervous system revealed by tyrosinated α-tubulin immunolabeling;c, Scanning electron microscopy (SEM) imaging of nerve net in a tentacle pocket (scale:15μm).d, Locomotory ciliated combs (SEM, scale:100μm).e, Glue-secreting cells – Colloblasts in tentacles (SEM, scale:50μm).f, Relationships among major animal clades with Choanoflagellates sister to all Metazoa.

Ctenophore Phylogeny

Although, relationships among basal animal lineages are controversial1,10–16, our analyses (Supplementary_Information_SD4) with Ctenophora represented byPleurobrachia andMnemiopsis suggest the placement of Ctenophora as the basal animal lineage (Fig. 1,Extended_Data_Fig. 3). Porifera was recovered sister to remaining metazoans (bs=100%) with Cnidaria sister to Bilateria (bs=100%,Fig. 1f). Shimodaira-Hasegawa (SH)-tests17(corresponding toExtended_Data_Fig. 3a, b, c with 586 gene matrix), rejected both Eumetazoa (sponges sister to all other metazoans) and Coelenterata (Cnidaria+Ctenophora). Placement of Ctenophora at the base of Metazoa also provides the most parsimonious explanation of the pattern of global gene gain/loss seen across major animal clades (Fig. 3,Supplementary_Table_14a, bS). Transcriptome data from ten additional ctenophores (Supplementary_Table_13S) with stricter criteria for orthology inference (Supplementary_Methods SM7), also placed ctenophores basal, albeit with less support (Extended_Data_Fig. 3d). When the most conserved set of genes was considered (Supplementary_Information_SM7.5/SD4.3), the topology was unresolved. Weak support is likely due to underrepresentation of comparable transcriptomes from sponges and large protein divergence. Nevertheless, SH-tests based on expanded ctenophore sampling (with a reduced 114 gene matrix due to lack of other ctenophore and sponge genomes –Supplementary_Methods_SD7.2) also rejected Coelenterata but not Eumetazoa. Importantly, relationships within Ctenophora were strongly supported (Fig. 2). Both cydippid and lobate ctenophores, previously viewed as monophyletic clades, were recovered as polyphyletic, suggesting independent loss of both the cydippid larval stage and tentacle apparatus. Interestingly, Platyctenida was the second basal-most branch in the Ctenophore clade, suggesting their benthic and bilaterial nature are secondarily derived.

Figure 3. Gene gain and gene loss in ctenophores.

Figure 3

a, Predicted scope of gene loss (blue numbers – e.g.–4,952 in Placozoa) from the common metazoan ancestor. Red and green numbers indicate genes shared between bilaterians and ctenophores (7,771), as well as between ctenophores and other eukaryotic lineages sister to animals, respectively. Text on tree indicates emergence of complex animal traits and gene families.b, Uniquely shared and lineage-specific genes among basal metazoans. Values under species names indicate number of genes (*) without any recognized homologs (e-value is 10−4) vs the total number of predicted gene models in are relevant species (Supplementary Table_14bS).

Figure 2. Phylogenomic reconstruction among major ctenophore lineages.

Figure 2

Cydippid (Euplokamis,Pleurobrachia,Dryodora, andMertensiidae) and lobate (Beroe,Mnemiopsis andBolinopsis) ctenophores were polyphyletic, suggesting independent loss of both cydippid larval stage and tentacle apparatus as well as independent development of bilateral symmetry in benthic/aberrant ctenophores,Vallicula andCoeloplana (Supplementary_Data SD4).

A highly reduced complement of animal-specific genes is a feature shared for the entire ctenophore lineage (Fig. 3,Supplementary_Table_15S). HOX genes involved in anterio-posterior patterning of body axes and present in all metazoans are absent in ctenophores and sponges18 (Supplementary Tables_17–18S). Likewise, canonical microRNA machinery (i.e.Drosha/Pasha,Supplementary Table_19S) is lacking inPleurobrachia and other ctenophores. Using small RNA sequencing fromPleurobrachia, Bolinopsis andBeroe, we were unable to experimentally detect microRNAs (Supplementary_Data_SD5.4).Pleurobrachia also lacks major elements of initiate innate immunity such as pattern recognition receptors (Toll-like, Nod-like, RIG-like, Ig-TIR) and immune mediators, MyD88 and RHD TFs, that are present in bilaterians, cnidarians and, in divergent forms, in sponges19,20 (Supplementary_Table_20S).

Key bilaterian myogenic/mesoderm-specification genes are absent inPleurobrachia’s genome and transcriptomes of ten other ctenophores (Supplementary_Tables_35S). These data suggest that muscles21 and, possibly, mesoderm evolved independently in Ctenophora to control the hydroskeleton, body shape and food capture. Thus, ctenophores might have independently developed complex phenotypic plasticity and tissue organization, raising questions about the nature of ctenophore-specific traits such as their unique development, combs, tentacles and aboral/apical organs, nervous systems.

Ctenophore Innovations

To assess genomic bases of ctenophore-specific innovations, we performed RNA-seq profiling of the major developmental stages (Fig. 4a, b) as well as adult organs and identified genes differentially expressed in these structures. ManyPleurobrachia genes, that have no homologs in other species, are specifically expressed and most abundant during the 4- to 32-cell cleavage stages as well as in tentacles, combs and the aboral organ (Fig. 4b,Extended_Data_Fig. 4). Thus, structures that are known as ctenophore innovations (Fig. 1d, e) have the largest complement of highly expressedPleurobrachia/ctenophore specific-genes. These data suggest extensive gene gain in cell lineages associated with early segregation of developmental potential leading to ctenophore-specific traits in structures controlling feeding, locomotion and integrative functions; a finding consistent with hypothesized ‘orphan’ genes contributing to variation in early development and evolution of novelties22,23.

Figure 4. Nature of Ctenophore Innovations.

Figure 4

a, Main developmental stages inPleurobrachia from eggs to the cleavage (2–64 cells), gastrulation (1–3 hrs) and formation of cydippid larvae (~24 hours).b, Hierarchical clustering of approximately 400 ctenophore-specific genes differentially expressed among different development stages and adult structures as revealed by RNA-seq experiments. Color index as follows: black indicates highest level of expression, followed by purple, red then down to white indicating no expression. Most of these ctenophore-specific genes are primarily expressed during 4–32 cell stages (asterisks). The red circle indicates a subset of novel genes uniquely expressed in combs, tentacles and the aboral organ. These genes lack recognized homologs in other organisms.c, Diversity and differential expression of RNA editing genes inPleurobrachia development and adult tissues (RNA-seq). ADAR1 has highest expression level in early cleavage stages while ADAR2-3 and ADAT1-2 are most abundant in the combs.d, Morphological appearance of neurons during 3rd day of development (the top insert, neuronal cell bodies are stained tyrosinated α-tubulin antibodies, red arrows) correlates with abundant expression of multiple iGluR receptors suggesting that Glutamate plays an important role as an intercellular messenger. Muscles formed well before neuronal differentiation at end of 1st day of development (the bottom insert, phalloidin staining, yellow arrow); white arrow points to the embryonic mouth with hundreds of cilia inside. Inc andd expression levels of RNA editing or iGluRs genes shown as a normalized frequency of sequence reads for a given transcript from all RNA-seq data for each developmental stage (Supplementary Methods).

Examples of known metazoan gene families that are considerably expanded in Ctenophora (Supplementary_Data_SD5, Table_16S), include collagens, RNA editing enzymes and RNA-binding proteins (Supplementary_Data_SD5).Pleurobrachia’s genome encodes the most RNA editing enzymes (ADAR1-4/ADAT1-3/CDA1-2) reported among metazoans24,25 (Supplementary_Data_SD5.5), possibly acting as the generalized mechanism generating posttranscriptional diversity and ctenophore-specific traits in locomotory and integrative structures (combs+aboral organ). Matching expansion of RNA regulatory mechanisms,Pleurobrachia has more RNA binding proteins (RBPs, especially RRM/ELAV, KH and NOVAs26,27,Supplementary_Table_21S) than any basal metazoan or choanoflagellate examined. Dozens of RBPs are selectively expressed and abundant during 8–64 cell stages (Supplementary_Table_31S), and might contribute to sequestration of RNAs and segregation in developmental potential leading to early cell-fate specification.

Phenotypic complexity positively correlates with presence and high differential expression of 92 homeodomainPleurobrachia genes (Supplementary_Data_SD5.2 and Table_17S); 76 genes reported inMnemiopsis18, whereas theAmphimedon homeodomain complement consists of only 32 genes. In contrast, some developmental pathways are either absent (Hedgehog, JAK/STAT) or have reduced representation in ctenophores (TGF-β, Wnt, Notch). Surprisingly, mostWnts are weakly expressed duringPleurobrachia development, while the ctenophore-specific subtypeWntX is primarily restricted to adult neuroid elements such as polar fields, aboral organ and tentacular conductive tracts (Extended_Data_Fig. 5e) suggesting a distinct molecular makeup neural systems.

Parallel Evolution of Neural Organization

Extensive parallel evolution of neural organization in ctenophores is the most evident. Compared to other animals with nervous systems, many genes controlling neuronal fate and patterning (e.g. Neurogenins/NeuroD/Achaete-scute/REST/HOX/otx) are absent in the ctenophores we sampled. Orthologs of pre-and postsynaptic genes also have limited representation (Supplementary_Table_34S), and they lack components (e.g. Neuroligin) critical for synaptic function in other eumetazoans.

Importantly, our combined molecular, ultra-sensitive metabolomic, immunohistochemical and pharmacological data strongly suggest ctenophores do not use serotonin, acetylcholine, dopamine, noradrenaline, adrenaline, octopamine, histamine or glycine as intercellular messengers (Extended_Data_Fig. 6,7g,Supplementary_Data_SD5.8, Tables_22–26S). Lack of ionotropic receptors for these molecules in ctenophores is consistent with this conclusion (Supplementary_Table_26aS). Most synthetic genes for neurotransmitter pathways are absent in non-metazoan opisthokontsMonosiga andCapsaspora suggesting they are cnidarian/bilaterian innovations.

But, what are the ctenophore transmitters? Physiological and pharmacological tests suggest that L-glutamate is a candidate neuromuscular transmitter inPleurobrachia (Fig. 5b,Extended_Data_Fig. 7), able to induce rapid inward currents and raise intracellular Ca2+ in muscle cells causing muscle contractions at nanomolar concentrations (10−7M). In contrast, all other classical neurotransmitters were ineffective even in concentrations up to 5×10−3M while D-glutamate as well as L-/D-aspartate have significantly less affinity in these assays (Fig. 5b).

Figure 5. Emergence of Neural Organization inPleurobrachia.

Figure 5

a, Two neural nets inPleurobrachia as revealed by tyrosinated α-tubulin immunostaining. Top image shows subepithelial net with concentrations of neuronal elements in the polar fields and ciliated furrows, known as structures involved in sensory and motor functions respectively (blue arrow in right insert indicates location of a neuronal somata with individual neurites marked by red arrows). The bottom image shows neurons of mesogloeal net (arrows are neuronal somatas; arrowheads are neuronal processes). Note, phalloidin (a muscle marker) did not stain these cells. Scale: 120μm (top); 10μm (bottom images).b, L- glutamate (10−7–10−3 M) induced action potentials in muscle cells whereas other transmitter candidates were ineffective even at concentrations up to 5mM. Typical responses of ctenophore muscle cells to local pulses of a transmitter application were recorded both as individual action potentials (whole-cell current-clamp mode) and video contractions from a single muscle cell. The graph shows normalized responses from the same muscle cell indicating L-glutamate is the most potential excitatory molecule compared to D-glutamate or L-/D-aspartate (Supplementary Methods).c, Key molecular innovations underlying neural organization in ctenophores. Bars indicate presence or relative expansions of selected gene families in all basal metazoan lineages from the inferred urmetazoan ancestor. The data suggests that sponges and placozoan never developed neural systems, or, assuming that pre-neuronal organization in the urmetazoan ancestor, sponges and placozoans lost their nervous systems. Either hypothesis point toward extensive parallel evolution of neural systems in ctenophores vs the Bilateria+Cnidaria clade.d, The aboral organ has the greatest diversity and highest expression levels of 12 gap junction proteins suggesting unmatched expansion of electrical signalling in this complex integrative organ - an analog of an elementary brain in ctenophores. Expression of different innexins shown as a summation of normalized frequencies of respective sequencing reads in RNA-seq data obtained from each developmental stage and adult tissues (Supplementary Methods).

The hypothesized role of glutamate as a signal molecules in ctenophores is supported by an unprecedented diversity of ionotropic glutamate receptors, iGluRs (Extended_Data_Fig. 7a, b,Supplementary_Table27S) – far exceeding the number of genes encoding iGluRs in other basal metazoans28. iGluRs might have undergone a substantial adaptive radiation in Ctenophora as evidenced by unique exon/intron organization for many subtypes and ctenophoran iGluRs form a distinct clade within the gene tree. Interestingly, during development,Pleurobrachia’s neurons are formed two days after the initial muscle formation, and first neurogenesis events correlate with co-expression of all iGlu receptors in hatching larvae (Fig. 4d). All cloned iGluRs also show remarkable cell-type specific distribution with predominant expression in tentacles, followed by combs and the aboral organ, revealing well-developed Glutamate-signalling in adults (Extended_Data_Fig. 7b). Additionally,Pleurobrachia contains more genes for glutamate synthesis (8 glutaminases) and transport (8 sialins) than any other metazoan investigated29,30. Although we detected Gamma-Aminobutyric acid (GABA,Supplementary_Tables 22–24S, and its localization in muscles), lack of pharmacological effects of GABA onPleurobrachia behavior and major motor systems, such as cilia, muscle and colloblasts, suggest that GABA is a by-product of glutamate metabolism by L-glutamic acid decarboxylase.

The first nervous systems are suggested to be primarily peptidergic in nature7. Although we did not find any previously identified neuropeptide homolog, secretory peptide prohormone processing genes (Supplementary_Table 31S) are present. We predicted 72 novel putative prohormones inPleurobrachia and found >50 homologs in other sequenced ctenophores (Extended_Data_Fig. 8,Supplementary Tables_28S,32S). Functions of these prohormone-derived peptides could include cell to cell signalling, toxins or involvement in innate immunity, or a combination. Several ctenophore-specific precursors are expressed in polarized cells around the mouth, tentacles and polar fields, suggesting a signalling role (Extended_Data Fig. 8b). They may be natural ligands for >100 orphan neuropeptide-like G-protein-coupled receptors31 identified inPleurobrachia (Supplementary_Table_26b). A second example of neuropeptide receptor candidates is amiloride-sensitive sodium channels (ASIC), which are also known to be regulated by different classes of short peptides and protons32.Pleurobrachia’s genome has 29 genes encoding ASICs -more than any organism sequenced so far, and expression of most correlated with developmental appearance of neurons (Supplementary_Table_31S). ASIC expression is most abundant in tentacles, combs and aboral organs –structures enriched in neural elements and under complex synaptic control.

Moreover, ctenophores evolved an enormous diversity of electrical synapses (absent inNematostella, Amphimedon andTrichoplax) with 12 gap junction proteins (pannexins/innexins33 but not chordate-specific connexins) found inPleurobrachia. All pannexins/innexins have their highest expression in the aboral organ followed by tentacles and combs (Fig. 5d). The aboral organ, combs and tentacles have a relatively large diversity of ion channels (Extended_Data_Fig. 9b), confirming complex regulation of excitability in these structures. Non-metazoans lack pannexin orthologs suggesting that these are metazoan innovations with profound expansion of this family in ctenophores. However, the overall complement of voltage gated ion channels in ctenophores is reduced compared to other eumetazoans34 (Extended_Data_Fig. 9a).

Our genome-wide survey also indicates that some bilaterian and cnidarian pan-neural markers are present (e.g. 3elav andmusashi genes), but they are not expressed in neurons; a finding consistent with early divergence and extreme parallel evolution of neural systems in this lineage (Extended_Data_Figs.5,9b).

Discussion

Figure 5c summarizes key molecular innovations underlying neural organization in ctenophores. Evidently, with an astonishing different molecular and genomic makeup, ctenophores have achieved complex phenotypic plasticity and tissue organization. Thus, ctenophores might represent remarkable examples of convergent evolution including the emergence of neuro-muscular organization from the metazoan common ancestor without differentiated nervous system orbona fide neurons (Extended_Data_Fig. 10b,Supplementary_Data_SD15). The alternative “single-origin-hypothesis”, where the common ancestor of all metazoans had a nervous system with complex molecular and transmitter organization including all classical cnidarian/bilaterian transmitters and neurogenic genes (Extended_Data_Fig. 10a), as a less parsimonious scenario. This hypothesis implies that ctenophores, despite being active predators, underwent massive loss of neuronal and signalling toolkits and then replaced them with novel neurogenic and signalling molecules and receptors.

These findings might have implementations for regenerative and synthetic biology in designing novel signaling pathways and systems. In this case, ctenophores (‘aliens of the sea’) and their genomes present matchless examples of “experiments” in nature and the possible preservation of ancient molecular toolkits lost in other animal lineages.

ONLINE METHODS

Source material

Animals (Pleurobrachia bachei,Euplokamis dunlapae,Dryodora glandiformis, Beroe abyssicola, Bolinopsis infundibulum andMertensiidae sp) were collected at Friday Harbor Laboratories (Pacific North-Western Coast of USA) and maintained in running seawater for up to two weeks. Other species were collected at the Atlantic coast of Florida and around of Woods Hole, Massachusetts (Pleurobrachia pileus,Pleurobrachia sp.,Mnemiopsis leidyi) as well as central Pacific (Palau, Hawaii,Coeloplana astericola, Vallicula multiformis). Animals were anesthetized in 60% (volume/body weight) isotonic MgCl2 (337mM). Specific tissues were surgically removed with sterile fine forceps and scissors and processed for DNA/RNA isolations as well as metabolomics or pharmacological/electrophysiological tests. Whole animals were used for allin situ hybridization and immunohistochemical tests as described35. Genomic DNA (gDNA) was isolated using Genomic-tip (QIAGEN, CA) and total RNA was extracted using RNAqueous-Micro (Ambion/Life Technology, TX) or RNAqueous according to manufacturers’ recommendations. Quality and quantity of gDNA was analyzed on a Qubit2.0 Fluorometer (Life Technologies) and for RNA we used a 2100 Bioanalyzer™ (Agilent Technologies, CA). For all details seeSupplementary Methods sections S1.1–1.3.

Genome sequencing

All genomic sequence data forde novo assembly were generated on Roche 454 Titanium and Illumina Genome Analyzer IIx, HiSeq2000 and MiSeq instruments using both shotgun pair-end and mate-pair sequencing libraries with 3–9 kb inserts as summarized inSupplementary Tables S1–2. Shotgun sequencing was performed from a single individual. Due to a limited amount of starting gDNA, mate pair libraries were constructed from 10–12 individuals. In total, the genome sequencing is composed of 106, 568, 866, 588 bp or ~106.5 Gb of data, which corresponds to 590-665x physical coverage of thePleurobrachia genome (the size of theP. bachei genome is estimated to be ~160–180 Mb); seeSupplementary Methods sections S1.4–2.1.2.

Genome assemblies

ThePleurobrachia bachei draft genome was assembled using a custom approach designed to leverage the individual strengths of three popularde novo assembly packages and strategies: Velvet36, SOAPdenovo37, and pseudo-454 hybrid assembly with ABySS38. First, using filtered and corrected data, we performed individual assemblies from 454 and Illumina reads by the Newbler (Roche, Inc.) software. Then the merged/hybrid assembly was achieved using three individual assemblies (SOAPdenovo, Velvet, and ABySS/Newbler as described in theSupplementary Methods S2.2). Three gene model predictions were performed by Augustus39 and Fgenesh predictions with the Softberry Inc. Fgenesh++ pipeline40,41 to incorporate information from full-length cDNA alignments and similar proteins from the eukaryotic section of the NCBI NR database42. After initial gene predictions in each of the three sets of genomic scaffolds, we screened each set of gene models for internal redundancy with the BLASTP program from NCBI’s BLAST+ software suite43. A model was considered redundant if it: had 90% identity to other model; the alignment between the two models had a bit score of at least 100 and the model was shorter than the other model.

Scaffolds producing these gene models were pooled and then screened for prokaryotic contamination using UCSC’s BLAT software package44 to produce the draft genome assembly version 1.0 (statistics can be found inSupplemental Table 5S andSupplementary Methods S.2).

Genome annotation

For annotation, gene models were uploaded to the In-VIGO BLAST interface, a blastp alignment of gene models was performed against the entirety of NCBI’s non-redundant protein database and the Swiss-Prot protein database, and subsequently annotated in terms of Gene Ontology and KEGG pathways as well as Pfam domain identification. Transposable elements (TEs) were identified using not only WU-BLAST and its implementation in CENSOR but also databases for all known classes, superfamilies and clades of TEs described in the literature and/or collected in Repbase45. Detected sequences have been clustered based on their pairwise identities by using BLASTclust. All autonomous non-LTR retrotransposons have been classified based on RTclass146. To merge partially predicted, non-redundant gene models with assembled transcriptome data, a custom Java tool was developed. This Java tool extended partial gene model predictions based on using transcriptome sequences to bridge 5′ and 3′ fragments of partially predicted genes. Using this Java tool, analysis of alignments of non-redundant gene models to assembledPleurobrachia transcriptomes resulted to 19,523 (Supplementary Table 26S) gene models. These gene models were employed to also identify their possible homologs in assembled transcriptomes from 10 other ctenophore species sequenced (Supplementary Tables 10S and 11S). All genomic sequences were submitted to NCBI on SRA accession number Project: SRP001155 (Supplementary Methods S.3.1–3.2.)

Transcriptome sequencing and annotation

Three sequencing technology platforms were used for transcriptome profiling (RNA-seq): Roche 454 Titanium, Illumina HiSeq2000 and Ion Proton/PGM (Ion Torrent, Life Technologies). RNA-seq was performed from all major embryonic and developmental stages (1-cell, 2-cells, 4-cells, 8-cells, 16-cells, 32-cells, 64-cells, early and later gastrula, 1-day, and 3-days larvae), major adult tissues and organs (combs, mouth, tentacles, stomach, the aboral organ, body walls), and whole body ofPleurobrachia bachei. We developed a reduced representation sequencing protocol for the 454 andIon Torrent sequencing platforms that can detect low abundance transcripts47. The method reduces the amount of sequencing and gives more accurate quantification and additional details of the procedure are reported elsewhere47,48. In summary, we have generated 499,699,347 Reads or ~47.9 Gbp to achieve approximately 2,000x coverage of thePleurobrachia transcriptome.

In addition, Illumina HiSeq sequencing was also performed with RNA extracted from the following ctenophore species:Euplokamis dunlapae,Coeloplana astericola, Vallicula multiformis, Pleurobrachia pileus,Pleurobrachia sp. (collected from the Middle Atlantic and later identified as a subspecies ofP. pileus),Dryodora glandiformis, Beroe abyssicola, Mnemiopsis leidyi, Bolinopsis infundibulum, and an undescribed species which belongs to the family Mertensiidae;Supplementary Table 3S). Each sequencing project was individually assembled using the Trinityde novo assembly package49 and in selected cases using MIRA. Reads from developmental stages were also assembled using the CLCBio Genomics Workbench. Prior to each assembly, reads were quality trimmed and had adapter contamination removed with cutadapt50. Full summaries of the transcriptome assemblies are presented inSupplementary Table 4S and 10S. Each transcriptome was mapped to thePleurobrachia genome, and aligned to both NCBI’s non-redundant protein database (NR) and the UniProtKB/Swiss-Prot (SP) protein database. Gene Ontology51 and Kyoto Encyclopedia of Genes and Genomes52,53 (KEGG) terms were associated with each transcript. By first translating transcripts in all six reading frames, Pfam/SMART domains54 were assigned to each reference transcriptome.

Each reference transcriptome and its full set of annotation and expression data was uploaded to our transcriptome databasehttp://moroz.hpc.ufl.edu/slimebase2/browse.php for downstream analysis and visualization55,56. The database is integrated with UCSC type genome browser. Via the genome project homepagehttp://moroz.hpc.ufl.edu/ all datasets have direct download options. Quantification of gene expression profiling was performed on all transcriptional data as described insupplementary methods S4.4). Hierarchical clustering was performed by Spotfire agglomerative algorithm. All primary transcriptome data was submitted to NCBI on SRA accession number Project: SRP000992. (Details seeSupplementary Methods S4.1–4.2.3).

Phylogenetic analyses

To reconstruct basal metazoan phylogeny (see controversies in10–15,57), we conducted two sets of phylogenomic analyses using tools described elsewhere58. All analyses included new data fromPleurobrachia bachei and the spongesSycon (Calcarea) andAphrocallistes (Hexactinellida). For the first set of analyses, Ctenophora was represented by two species ofPleurobrachia andMnemiopsis leidyi. Initial analyses included the taxa inSupplementary Table 12S. For a subsequent analysis, sampling within Ctenophora was expanded to include ten additional taxa, each represented by a relatively deeply sequencedIllumina transcriptome (Supplementary Table 13S). In order to reduce noise in the phylogenetic signal, we employed strict criteria to exclude paralogs, highly derived sequences, mistranslated sequence regions, and ambiguously aligned positions in sequence alignments. Analyses were conducted in RAxML 7.2.759,88 using maximum likelihood (ML) with the CAT +WAG + F model. Topological robustness (i.e., nodal support) for all ML analyses was assessed with 100 replicates of nonparametric bootstrapping. Details of phylogenomic analyses are presented inSupplementary Methods S7. SH-test89 as implemented in RAxML with the PROTGAMMAWAGF model17.

In order to examine evolution of single genes or gene families, alignments were performed with either ClustalX260–62 or Muscle63 then, if appropriate, either trimmed manually or trimmed using GBlocks64 to exclude ambiguously aligned positions. Once alignments were obtained, gene trees were reconstructed in MEGA 565 using ML with the Whelan and Goldman (WAG) model. The bootstrap consensus tree was inferred from 100 replicates. All positions containing gaps and missing data were eliminated. Pfam composition54, Gene Ontology51, and KEGG52,53 were used to further validateP. bachei orthologs. Analyses of gene gain and gene loss were performed using custom scripts as described elsewhere66 and inSupplementary Methods S7.

Analysis of DNA methylation

ELIZA based colorimetric assays (Epigenteck, NY) were performed to quantify both global 5-mC and 5-hmC methylation in theP. bachei genome. A total of 6 individualP. bachei and three Rat (positive control) were used (Supplementary Methods S1.2). Three biological and technical replicates were performed for every sample. Absolute quantification of 5-mC and 2hmC were determined and date is reported as an mean ± S.E.M (Supplementary Methods S8).

Molecular cloning,in situ hybridization and immunohistochemistry

Methods were similar as reported elsewhere35,47,48,67 with some modifications (Supplementary Methods S9–S11).

Scanning electron microscopy

Animals were fixed in 2.5% glutaraldehyde in 0.2 M phosphate-buffered saline (pH=7.6) for 3–4 hours at room temperature, and washed. For secondary fixation, we used 2% osmium tetroxide in 1.25% Sodium Bicarbonate for 2–3 hours at room temperature. After dehydration in ethanol samples were was placed for drying in Samdri-790 Critical Point Drying. After drying the samples were coated on Sputter Coater. SEM observations and recordings were done on NeuScope JCM-5000 microscope (Supplementary Methods S12).

Electrophysiological methods, calcium imaging and pharmacological assays

Patch electrodes for extracellular and whole-cell recordings were pulled from borosilicate capillary (P-87, Sutter Instruments). All currents were recorded using an Axopatch or 200B amplifier controlled by a Digidata 1322A and pClamp 9.2. Action potentials (APs, spikes) were recorded in track mode using cell-attached loose-patch configuration. Whole-cell currents were recorded in voltage clamp mode at a holding potential of −70 mV. Neurotransmitter candidate (seeSupplementary Method S15) application for both extracellular AP and whole cell recordings were performed with a rapid solution changer, RSC-160 (Bio-Logic-Science Instruments, France). Data were analyzed with Clampfit 9.0 (Molecular Devices) in combination with SigmaPlot 10.0. Videomicroscopy and time-lapse series were acquired with QImaging EXi CCD camera using DIC mode of Nikon Eclipse 2000 inverted microscope. Calcium imaging was performed on isolated ctenophore muscle cells using Olympus IX-71inverted microscope equipped with a cooled CCD camera (ORCA R2, Hamamatsu). Cells were injected with calcium sensitive probe (Fluo-4, ~5μM) through patch pipette. Fluorescence imaging was performed under the control of Imaging Workbench 6 software. Stored time series image stacks were analyzed off-line using Imaging Workbench 6, Clampfit 10.3, SigmaPlot 10/11 or exported as TIFF files into ImageJ 1.42. Pharmacological tests and behavioral assays with video recording were performed on intact animals in 5–40 L aquaria or on semi-intact preparations in a Sylgard-coated Petri dish with free cilia beating and muscle contractions. To monitor and quantify cilia movements we used glass microelectrodes filled with 2M potassium acetate with resistances of 5–20 MΩ with electrical signals recorded by A-M System amplifiers (Neuroprobe 1600) and Gould Recorder (WindoGraf 980).

Determination of the presence of classical neurotransmitters by capillary electrophoresis (CE)

Two CE separation techniques were employed to analyze tissue extracts for the presence of a number of neurotransmitters (Supplementary Table 18S). While both methods used CE separations, complimentary detection methods, laser-induced native fluorescence (LINF)68 and electrospray ionization mass spectrometry (ESI-MS)69,70, were used to ensure broad coverage and low detection limits for the specific analytes of interest. Whole body of small animals as well as individual organs and tissues were removed, rinsed with ultrapure water and analytes were extracted using 49.5/49.5/1, methanol (LC-MS grade)/water/glacial acetic acid (99%) by volume, homogenized, centrifuged and supernatant was removed and frozen at −80°C until analysis. The CE-LINF instrument employed ultraviolet excitation at 264 nm and the native fluorescence emission collected and recorded using a UV-enhanced CCD array (Spec-10; 2KBUV/LN; Princeton Instruments; Trenton, NJ, USA). CE separations were performed by hydrodynamic injection of 10 nL of sample and using 25 mM citric acid (pH 2.5, applied voltage +30 kV) or 50 mM borate (pH 9.5, applied voltage +21 kV). Analytes were identified based on comparison of both the migration time and fluorescence spectrum to that of standard mixtures of analytes. CE-ESI-MS analysis was performed using a Bruker Microtof or a Maxis (Bruker Daltonics; MA) mass spectrometer for detection. All separations were performed using 1% formic acid in water as the electrolyte and applied voltage of +30 kV. Sheath liquid was 0.1% formic acid in 50/50 methanol/water. Samples were hydrodynamically injected for a total volume of ~ 6 nL. Mass spectra were collected and recorded at a rate of 2 Hz with calibration was performed using sodium formate clusters. Analytes were identified based on comparison of both the CE migration time and mass match to that of standard mixtures of analytes.

Extended Data

Extended Data Figure 1.

Extended Data Figure 1

a–e, Anatomy of the ctenophore,Pleurobrachia bachei A. Agassiz, 1860. Natural coloration of the major organs in live animal are shown.a, Details of the transparentPleurobrachia body are shown including,b, the pharynx and tentacle sheaths (pockets). Eight rows of comb plates, called ctenes, are made of giant compound cilia that diffract light – creating iridescence.c, Combs rows inPleurobrachia are constantly beating. The mouth and the aboral organ (AO) are located at the opposite poles of the animal (a, c). The AO controls complex coordinated behaviors of the animal;d, Ciliated furrows connect the AO and the ctenes to mediate behavior.e, Tentacles have numerous contractile tentillae used to capture food with specialized glue cells or colloblasts (See alsoFig. 1e of the main text).

f–h,Pleurobrachia neural nets and muscles. f, Comb plate muscles (red) were revealed usingin situ hybridization for β-tubulin and subepithelial neural net (green) revealed by tyrosinated α-tubulin immunostaining.g, In this image comb cilia (green) were stained using tyrosinated α-tubulin antibodies (green) where as underlying comb plate muscles were visualized by phalloidin (a muscle marker) that did not stain neurons.h, Organization of the subepithelial neural net around the Mouth as revealed by tyrosinated α-tubulin antibodies (whole mount preparation). Scale: 120 μm (f); 100 μm (g); 200 μm (h). SeeSupplementary Methods SM10 and SM11.

Extended Data Figure 2. DNA methylation and active DNA demethylation inPleurobrachia bachei.

Extended Data Figure 2

CpG DNA methylation facilitates the elimination of CpG dinucleotides over evolutionary time66a, Histogram shows relative occurrences of different dinucleotides in genomes ofP. bachei (red bars),Drosophila melanogaster (green bars, no DNA methylation) andHomo sapiens (blue bars). TheP. bachei genome contains 2.3 % CpG dinucleotides, which is much lower than the expected random frequency and, therefore, indicative of a genome that undergoes methylation compared to humans66.b,DNMT genealogy tree. The enzyme DNA methyltransferase (DNMT), which catalyzes transfer of a methyl group to DNA to form 5- methyl cytosine (5-mC)147, is present inPleurobrachia.c,TET family of enzymes catalyzes active DNA demethylation via formation of 5-hydroxymethyl cytosine (5-hmC, the 6th DNA base). RNA-seq profiling reveals differential expression for DNMT and TET-like genes during development and in adultP. bachei. Both DNMT and TET-like genes are predominantly expressed during cleavage starting from the 1st division. However, the TET-like gene is also highly expressed in adult combs (asterisk). Y-axis shows a normalized expression level for each transcript.d, ELIZA based colorimetric assays validate the presence of both 5-mC and 5-hmC in theP. bachei genome (the rat brain is used as a positive control; n=6 forPleurobrachia and n=3 for rat; data shown as mean ± s.e.m, see theSupplementary Methods SM8 andSupplementary Data section SD3 for details).

Extended Data Figure 3.

Extended Data Figure 3

a, Phylogeny of Metazoa based on 586 genes. Topology inferred using RAxML 7.2.7 and maximum likelihood (ML) with the CAT +WAG + F model with all taxa from theSupplementary Table 12S. Bootstrap support values are listed at each node.Color coding:purple –Ctenophorayellow – Porifera,pink – Cnidaria,light blue – Bilateria.b,Removal of fast-evolving taxaTrichoplax andCaenorhabditis improves topological robustness. Topology inferred using RAxML 7.2.7 and maximum likelihood (ML) with the CAT +WAG + F model with all taxa fromSupplementary Table 12S exceptTrichoplax andCaenorhabditis. Bootstrap support values are listed at each node.c,Removal of distant out-groups such as Fungi and Filasterea further improves topological robustness. Topology inferred using RAxML 7.2.7 using maximum likelihood (ML) with the CAT +WAG + F model with all taxa fromSupplementary Table 12S exceptTrichoplax,Caenorhabditis, and non-choanoflagellate outgroups. Bootstrap support values are listed at each node.d,Analysis with improved ctenophore taxon sampling based on 114 genes. Topology inferred using RAxML 7.2.7 using maximum likelihood (ML) with the CAT +WAG + F model with all taxa fromSupplementary Table 13S. Bootstrap support values are listed at each node.

Extended Data Figure 4.

Extended Data Figure 4

a, Identification of tentacle-specific transcripts. The left photo shows scanning electron micrograph of aPleurobrachia tentacle with two branching tentillae densely covered with hundreds of colloblasts or glue cells. Comparative transcriptome (RNA-seq) profiling among major organs allowed us to identify several dozen genes differentially or uniquely expressed in tentacles. The histogram shows illustrative examples of some of these genes with a normalized expression level (Y-axis) for each represented transcript. One of thesePleurobrachia-specific genes we named Tentillin (green arrow).In situ hybridization experiments (n=9) revealed a remarkable cell-specificity expression patterns for Tentillin in all main tentacle branches and tentillae, possible labeling colloblasts or associated secretory cells.b,Identification of comb-specific transcripts. The left photo shows a microscopic image of one comb row from an intact animal. The natural coloration is a reflection of the beautiful iridescence patterns produced from large cilia forming combs. Comparative transcriptome (RNA-seq) profiling among major organs allowed us to identify several hundreds of genes differentially or uniquely expressed in combs. The histogram shows illustrative examples of some of these genes with a normalized expression level (Y-axis) for each represented transcript (seeSupplementary Methods S4.2.3.6, S4.2.3.7 and SM10, all sequences used in the analysis can be found inSupplementary Tables 29S,30S and32S).

Extended Data Figure 5.

Extended Data Figure 5

a, b,Dicer andArgonaut, are predominatly expressed in structures associated to sensory and integrative functions. These include the aboral organ, polar fields and combs. Note, a relatively weak staining of other cell types in the skin and following ciliated furrows inDicer andArgonaut preparations (top two images).c,d,Pleurobrachia ELAV is Expressed in Combs and not in neurons. ELAVs are RNA binding proteins and they are considered as pan-neuronal markers (seeSupplementary Data SD5.6.1). However, inPleurobrachia ELAVs’ expression has not been detected in neural tissues or cells with recognizable neuronal like appearances.In situ hybridization forPleurobrachia ELAV3 (c–d) shows the highest levels of expression in the adult comb plate but not in any of the neural tissues or organs enriched with neurons such as the aboral organ and polar fields.e,WntX is selectively expressed in the aboral organ (AO) and major conductive pathways ofPleurobrachia suggesting its involvements in integrative and neural-like functions (in situ hybridization on a whole-mount preparation). A. One of the highestWntX expressions is found in AO and ciliated furrows whereas the polar fields showed a moderate expression level associated to their central regions.In situ hybridization was performed on whole mounts using DIG labeled probes (see details in theSupplementary Methods, allin situ hybridization were performed at least on 4–5 different animals and these are representative photos for these experiments). Scales: 500 μm (a–d). 800 μm (e).

Extended Data Figure 6.

Extended Data Figure 6

a–c, Absence of Serotonin in ctenophores. Here, we used nanoliter volume sampling, capillary electrophoresis separation, and wavelength-resolved native fluorescence detection as described for ultra-sensitive assay of 5-hydroxytryptamine (serotonin or 5-HT) and related metabolites (a, the top electropherogram and the table with standards used). Limits of detection (LODs) range from the low attomole to the femtomole range, with 5-HT LODs being approximately 20–50 attomoles.b, Using this assay we failed to detect 5-HT inPleurobrachia (bottom left, n=6) butc, 5-HT was reliable detected in the hemichordateSaccoglossus (bottom right) and molluscs62. See details in theSupplementary Methods SM17 andSupplementary Table 22S for quantification.

Extended Data Figure 7.

Extended Data Figure 7

a, The Ionotropic glutamate receptors (iGluRs) are diverse and underwent substantial adaptive radiation within the Ctenophora lineage. Phylogenetic analysis showsPleurobrachia iGluRs share highest identity to each other forming a distinct branch on the tree topology (Supplementary Data SD5.9).b,Differential expressed of iGluR subtypes inPleurobrachia bachei (red and green labeling with fluorescentin situ hybridization protocols). Dark blue fluorescence is DAPI nuclear staining. Aboral organ –AO. Scale 100 μm (b1-2), 60 μm (b3), 50 μm (b4), 30 μm (b5), 200 μm (b6).c–f,Glutamate induced action potentials and currents in muscle cells. c, Typical responses of ctenophore muscle cells to glutamate pulses recorded extracellulary (as individual action potentials/contractions from a single muscle cell in response to local application of Glutamate, 1mM), andd, from the same cell in whole-cell current clamp mode with clear action potentials.e, Isolated muscle cell. Scale 25μm.f, Glutamate activated whole-cell currents recorded from the same cell (as inc). Time course of application is depicted by the diagram below the voltage signal. Two responses (inward current) are shown. The holding potential was −70mV (Supplementary Methods SM13-16).g, Representative electropherograms show capillary electrophoresis separation with laser induced fluorescence detection from different organs inPleurobrachia bachei (n=5) for transmitter candidate identification. The bottom electropherograms are standards (Supplementary Methods SM17 andSupplementary Tables 23–25S for quantification).

Extended Data Figure 8.

Extended Data Figure 8

a, Computational pipeline for prediction of secretory products inPleurobrachia and the overview of secretory products predicted from thePleurobrachia gene models (Supplementary Method and Data sections SM4.2.3.7 and SD5.11, respectively).b,Expression of novel secretory molecules in Ctenophores (DIG-labeledin situ hybridization,Supplementary Data SD5.11 andMethods SM 10). Each of predicted secretory prohormone was selected based upon its unique and/or highly differential expression pattern as revealed by RNA-seq profiling.Ctenophorin is uniquely expressed in polarized cells around the mouth ofPleurobrachia and we found its homologs in all ctenophore species we sequenced- here is its name.Tentillin is aPlerobrachia-specific gene, which is uniquely expressed in polarized secretory-like cells in tentillae and tentacles.Jansonin’s expression is primarily restricted to polarized cells located in the aboral organ and polar fields.b4, For comparison, we showed different but also cell-specific expression pattern of BarX Transcription Factor in cells of unknown identify localized in polar fields, comb plates and tentacles.c–d,Majority of predicted secretory products are expressed later in development and in adult organs ofPleurobrachia (RNA-seq).c, Expression patterns of 72 predicted prohormones inP. bachei indicates that 20 of them are present and differentially expressed in development (Supplementary Table 32S for allPleurobrachia precursor sequences). Surprisingly 5 of these precursor mRNAs were found starting from the 2nd cleavage stage whereas the rest are predominantly expressed on day 3 of development. This correlates to the first appearance of neurons inPleurobrachia cydippid larva (seeSupplementary Data SD5.11 andSupplementary Method section S4.2.3.6 for the RNA-seq analysis).

Extended Data Figure 9.

Extended Data Figure 9

a, Metazoan Ion Channel Complement. The 112 ion channels identified in thePleurobrachia genome are classified as voltage gated (v) or other gating such as second messengers. Receptors channels (R) are ligand-gated or ionotropic (iGluR, ChRN, HTR3, GABA and CLR) and indicated in the grey. Metazoan novelties indicate type of ion channels absent in the choanoflagellates, the sister group to all animals. Colored squares show channels: (i) primarily absent in Ctenophores (pink), (ii) secondarily lost in sponges or placozoans (dark yellow), (iii) eumetazoan (Cnidaria+Bilareria) innovations (blue), or (iv) examples of expansion of certain class of channels in some animal lineages (red). AllPleurobrachia sequences used in the analysis can be found inSupplementary Table 31S.b,Ion Channels are Predominantly Expressed in Tentacles, Combs and Aboral Organ. Hierarchical Clustering of 112 identified ion channels in developmental stages and adult tissues ofPleurobrachia. Adult organs involved in food capture and ciliated locomotion and integrative functions show significantly higher diversity and overall higher level of expression levels for most of ion channel types. Mobile tentacles had the highest expression of voltage gated channels, in particular Cav and Nav. The legend shows relative expression levels based on RNA-seq data (seeSupplementary Methods S4.2.3.6).

Extended Data Figure 10. Two alternative scenario of neuronal evolution.

Extended Data Figure 10

a,Single origin of the neural system (Monophyly) with possible loss of some neural molecular components in Ctenophores as well as the possible secondarily loss of the entire nervous systems in sponges and placozoans;

b,Multiple origins of neurons in animals as introduced and supported by this manuscript (see main text discussion section for details).

Supplementary Material

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supp table 31
supp table 32
supp table 33
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supp video1
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supp video2
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video legends

Acknowledgments

FHL for facilities during animal collection and Marine Genomics apprenticeships (L.L.M., B.J.S.); E. Dabe, G. Winters, J. Netherton, Caleb Bostwick for help with animal/tissue/RNA/DNA assays; Drs X-X Tan/F. Lu (SeqWright, Inc) and T. Tyazelova for sequencing. F. Nivens for videos. Supported by NSF (NSF-0744649/ NSF CNS-0821622 to L.L.M., NSF CHE-1111705 to J.V.S.), NIH (1R01GM097502,/R01MH097062, R21RR025699/ 5R21DA030118 to L.L.M., P30 DA018310 to J.V.S., R01 AG029360 to E.I.R.), NASA NNX13AJ31G (to K.M.H./L.L.M,/ K.M.K.), NSERC458115/211598 (/J.P.R.), University of Florida Opportunity Funds/McKnight Brain Research and Florida Biodiversity Institute (L.L.M.), Rostock Inc/A.V. Chikunov (E.I. R.); Grant from RF Government No 14.B25.31.0033 and NIH R01 AG029360 (E.I.R.). F.A.K./I.S.P/ R.D. were supported by HHMI (55007424), EMBO and MINECO (BFU2012-31329 and Sev-2012-0208). Contributions of AU Marine Biology Program_#117 and Molette Lab_#22.

Footnotes

Supplementary information is available in the online version of the paper.

Author Contribution: L.L.M. conceived the project, designed the experiments and wrote the manuscript. A.B.K., A.P.G., D.R., E.K., T.T., R.S.,T.P.M., E.I.R. and L.L.M. prepared gDNA, RNA samples and performed sequencing; I.S.P, F.A.K., V.V.S., F.Y., M.R.C., A.B.K., L.L.M. did assemblies, gene model prediction and annotations; K.M.K., K.M.H. performed phylogenomic analysis; A.P., A.B.K. and L.L.M. worked on gene family gain/loss analysis; F.A.K. and R.D. characterized protein divergence; S.D., C.D., J.V.S. and L.L.M. performed capillary electrophoresis/ microchemical metabolomic assays; A.P.G., A.B.K., E.B., E.I.R. did small RNA sequencing and analysis; K.B. and J.R. characterized immune gene complement; V.K. and J.J. characterized transposons, T.P.N and L.L.M. performed immunolabeling, electron microscopy and pharmacological assays; Y.B. and L.L.M. performed pharmacological, electrophysiological and imaging assays on muscles; D.O.G., M.R.C., A.B.K. and L.L.M. performed secretory peptide prediction; A.B.K. and L.L.M. analyzed RNA-seq data; A.B.K. performed methylation analysis; B.J.S., A.B.K. and L.L.M. analyzed developmental data; J.J.S., D.O.G., R.B., A.F., A.B.K. and L.L.M. performedin situ hybridization experiments; C.E.M. identified species and wrote their description and biology; all authors contributed to preparation the manuscript and the text.

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