Cnidarians are simple animals with radial symmetry that contain two layers of cells, ectoderm and endoderm. Mesoglea, a non-cellular matrix, is present between the two layers. Cnidarians are mostly predators but certain species may also scavenge dead animals or obtain nourishment from intracellular, photosynthetic unicellular algae, named zooxanthellae.

At least four toxic living classes of cnidarians are currently recognized by most systematists: Anthozoa, Hydrozoa, Scyphozoa and Cubozoa. Molecular phylogenetic methodologies based on DNA sequencing, allowed to determine that the Anthozoa are the basal group of cnidarians [1] (Figure 1). In fact, Anthozoa has a circular mitochondrial DNA, while Hydrozoa, Scyphozoa and Cubozoa have a linear molecule. Likewise the polyp preceded the medusoid form in the course of evolution [2].

There are two main types of life cycles in cnidarians. In Anthozoans, the polyp is the gamete-producing form and the cycle is embryo > larva > polyp. Medusozoans generally have an embryo > larva > polyp > medusa life cycle, in which the medusa is typically the sexual form. Figure 2 shows a typical life cycle of Anthoza [1].

Cnidaria feeding success relies on the presence of specialized poisonous cells, the nematocysts. These organisms have specialized subcellular organelles called cnidae with several structures and functions. Cnidae can be classified into three types: nematocysts, spirocysts, and ptychocysts. Nematocysts deliver the venom through the skin, whereas spirocysts are adhesive and ptychocysts are involved in protection. While Anthozoans have the three types of cnidae, medusozoans (Scyphozoans and Cubozoans) contain only nematocysts. The biological roles of toxins delivered by nematocysts include the capture and killing of prey, digestion, repelling of predators and intraspecies spatial competition [5]. Cnidarians are not just studied by their toxins and venoms, they are a source of marine natural compounds with therapeutically properties, namely antitumor activity [6]. Furthermore, voltage-gated ion channels toxins are studied as an inspiration for drugs design, not only therapeutic but also as insecticides [7].

The composition of cnidarian venoms is not known in detail, but they appear to contain a variety of proteinaceous (peptides, proteins, enzymes and proteinase inhibitors) and non-proteinaceous substances (purines, quaternary ammonium compounds, biogenic amines and betaines) [8]. As an example, palytoxin is a polyether from Palythoa, and caissarone is an iminopurine from Bunodosoma caissarum [9].

The venom is spread all over the body, in a mucous coat, that also protects them from predators, or it is located in the nematocysts. In a recent work, Moran and co-workers, reported that neurotoxin 1 from Nematostella vectensis is confined to ectodermal gland cells. Moreover, in Anthopleura elegantissima this toxin also appears in gland cells, whereas in Anemonia viridis is associated with both nematocytes and ectodermal gland cells [10]. Previously, Honma and co-workers also gave a hint for the same phenomenon when describing that gigantoxins were mostly derived from unknown organelles other than nematocysts [11]. Nematocysts are found mostly on the tentacles, but also exist in other organs such as in acrorhagi and acontia, particularly in certain species of the Actiniidae family, where they are used to fight with nonspecific non-clonemates or for purposes of defence or predation, respectively. Acrorhagi are located in a ring around the base of the tentacles (Figure 3a). Acontia are thin white or color threads attached at one end to the borders of the mesenteries. They can be protruded through the mouth, and in some cases through special pores (cinclides) in the body-wall, for purposes of defence or paralyses of prey (Figure 3b).

The Anthozoa class include sea anemones, and other anemone-like groups with skeletons (such as the “stony” scleractinian corals) and without skeletons (such as tube anemones), as well as sea pens, sea fans, blue corals, and black corals. The word Anthozoa comes from greek anthos, flower + zoon, animal, as sea anemones resemble flowers (Figure 3c).

Nematocysts possess a high concentration of polypeptides and proteins that act as neurotoxins, hemolysins and enzymes, which are responsible for a variety of harmful effects to humans. These toxins/venoms are only injected in the prey or predator after a mechanical or chemical stimulation [12]. In humans, toxins cause cardiotoxicity, dermatitis, local itching, swelling, erythema, paralysis, pain and necrosis [8]. In vivo effects of sea anemone toxins include neurotoxicity and cardiotoxicity.

Summarily, the cnidarians venom includes 3.5–6.5 kDa voltage-gated sodium (Na_V) channels toxins and 3–5 kDa voltage-gated potassium (K_V) channel toxins and ~20 kDa pore-forming toxins. The first type prevents inactivation of Na_V channels by stabilizing the open state conformations. This fact is due to the binding of the toxin to neurotoxin receptor site 3 [13]. K_V channel toxins reversible blocks potassium current and can block acid-sensing ion channels, which are permeable to several cations. The cardiotoxic effects of toxins includes arrhythmias, triggered by early after depolarizations resulting from incomplete Na_V channel inactivation, and systolic arrest due to myocardial cell calcium ion overloading [9].

Besides toxins, there are several other non-toxic proteins from sea anemones that are studied by its biological activities, such as fluorescent properties [14], but they will not be included in this review. However, we will discuss the importance of protease inhibitors as they adopt a structure that inhibits potassium channels.

In this review, we begin with a brief description of the Anthozoa phylogeny, followed by a general characterization of the sea anemone toxins and afterwards we focus on the major groups of toxins. We then refer to the state of the art techniques used for venom extraction. Afterwards we present the structure of the genes involved in toxin production and the three-dimensional (3D) structures of cnidarian toxins described to date. This review will be solely focused in the molecular diversity of sea anemone toxins. Other cnidarian toxins, as those from coral or jellyfish, will not be considered. More comprehensive information is available in a number of specific papers for jellyfish [15,16], cnidarians in general [2,5,17,18] and sea anemones [13,19,20].

Cnidarians are scattered around the world and have around 10,000 estimated species. The majority of the phylogenetic studies classified cnidarians based on morphological characters [21]. At the molecular level, the classification of cnidarians is not yet well established, namely for the order Actiniaria. The phylogeny of Actiniaria is at a suboptimal estimation level [3] and has been retrieved from the sequencing analyses of 12SrRNA, 16SrRNA, 18SrRNA, 28SrRNA and COIII genes [22,23,24,25]. As referred by Turk and Kem [2], the comprehension of the phylogenetic relationships among Anthozoa members will give insights into the evolution of theirs toxins. Thus, a review about sea anemone toxins could not be dissociated from the Anthozoa phylogenetic characterization.

Besides the few studies on the phylogeny of Actiniaria, some other studies have also been done on the population genetics of these animals. Nonetheless, the majority of those works focus on other Orders, especially on corals. Indeed, few studies were done at the intraspecific level on Actiniaria. Population genetics of Actinia spp. assessed with enzyme electrophoresis showed that Actinia nigropunctata from Madeira Island (Portugal) is in fact a different species from all the others in the study, as well as Actinia equina from Africa [26]. Darling and co-workers in 2006 studied the Nematostella vectensis introduced along the Pacific coast of North America and the southeast coast of England, using 10 polymorphic microsatellite loci, and find high variability from Hardy-Weinberg equilibrium as a result of population genetic structure and reproductive plasticity [27].

Considering the molecular markers surveyed in Cnidarians until now, the variation in mitochondrial Citochrome Oxidase I (COI), within and between species, is much lower in Anthozoa compared to Medusozoa. Low identification success and substantial overlap between intra- and interspecific COI distances render the Anthozoa unsuitable for DNA barcoding [28], with COI p-distances among Anthozoa species being equal to 1% [29]. Shearer and co-workers [30] showed that nuclear markers in Anthozoa have much higher substitution rates and therefore should be used instead of mitochondrial genes.

The reduce knowledge on sea anemones phylogeny make it difficult a direct comparison with the toxin genes phylogeny. While previous studies showed a reduced level of congruence between species phylogeny and the toxin gene phylogeny, further research is needed to better clarify this pattern. Such findings may not be unusual due to distinct patterns of toxin gene evolution (e.g., gene duplication/gene loss, horizontal gene transfer, and lineage sorting and diversification). However, future studies are needed to better elucidate the phenomena behind the acquisition and evolution of the toxin genes in Anthozoa.

Concerning the phylogeny of toxins, we assessed the phylogenetic relationships of Na_V channel and K_V channel toxins. In order to systematize the information, we have assessed a phylogenetic tree of cytolysins using only Actinoporins with evidence at transcript level and with full-length sequences. A multiple sequence alignment of amino acids with 533 sites, was made with WebPrank [31] followed by an analysis to choose the best fit model for protein evolution with ProtTest [31], that gave WAG model. A Maximum Likelihood tree reconstruction was made in Mega 5 [32] using 100 bootstrap inferences. A discrete Gamma distribution was used to model evolutionary rate differences among sites (4 categories), Figure 4. (The alignment is available upon request to the corresponding author.)

Considering the phylogenetic tree of cytolysis, two major groups can be defined; one including the proteins without the MACPF domain (I) and the other comprehending those with the MACPF domain (II). Within the major group “I” three clusters can be identified (III to V). Toxins from Actiniidae family members are clustered in group III. In Group IV cluster toxins from Stichodactilidae family members and toxins from Oulactis orientalis, (Actiniidae). In fact, toxins from Oulactis are more closely related to Stichodactilidae than to Actiniidae toxins. As mentioned previously, Or-A and Or-G and RTX-S-II and RTX-A from Hecteractis crispa have in common (albeit others characteristics), the substitution of a Trp by a Leu in the position Trp¹¹² of Equinatoxin-II. Moreover, the conserved RGD sequence that occurs in Sticholysin-II, RTX-A and Equinatoxin-II, in the toxins from Oulactis is replaced by the GGD sequence. The cluster V includes the Src-I and the toxins from Alisiidae family. The only member of Sagartiidae family (Src-I), has the EGD sequence instead of the RGD motif. Toxins of Alisiidae family members, share a similar gene organization with three exons (two introns). In addition the RGD motif is replaced by the KPS tripeptides in PsTX-20A and Avt.

Regarding the sea anemones phospholipases toxins, the study of Romero and co-workers [33] comparing PLA2 from Condylactis gigantea (Actiniidae family member), CgPLA2, with the other PLA2s from five animal phyla, suggested that sea anemones PLA2s form a monophyletic group. Within this group, CgPLA2 showed to be closer to the Adamsia carcinoapados (Hormathiidae family member) PLA2, AcPLA2, than to others of Nematostella vectensis, suggesting a significant divergence from the latter.

In the first decades of the 20th century, it was practically impossible to isolate and chemically characterize venom compounds, as the biochemical techniques for isolating such natural products hardly existed [2]. However, nowadays scientists developed several techniques to obtain the venom of particular structures such as acrorhagi or nematocysts, and to separate the venom into fractions. In this sense, there has been an increase in the number of publications on the subject of “cnidarian toxins”. Figure 5 shows the number of publications in Pubmed, retrieved using the query “Cnidaria toxins” on 5 March 2012. In fact, it is expectable that with the deep sequence platforms, much more data will become available at genomic level allowing to better understand the evolution of cnidarians toxins and the discover of the pharmaceutic and therapeutic properties of such compounds. Deep sequencing transcriptomics is the sequencing of the complete set of cellular transcripts at a specific stage or condition, and in that sense Johansen and co-workers [34] and Rodríguez and co-workers [35], pioneered the use of cDNA high-throughput sequencing with 454 pyrosequencing in the discovery of new toxins. The first publications on “cnidarian toxins” were about crude extracts (e.g., jellyfish crude extracts) and not about isolated toxins. In the following years, sea anemones toxins started to gain some relevance. A partial purification of a toxin from the tentacles of Condylactis gigantea was made by gel filtration [36]. Afterwards, three neurotoxic peptides were isolated from Anemonia viridis by cm-cellulose and sephadex chromatography [37]. The ATX-II amino acid sequence published by Wunderer and co-workers was the first cnidarian toxin to be determined [38]. At same time, in another laboratory, peptides from Anthopleura were also studied [2].

Voltage-gated ion channels underlie electrical excitability in cells, and they also play important roles in non-excitable cells. Voltage-gated channels open in response to changes in membrane potential, allowing ions to flow down the electrochemical gradient across the cell membrane, being thus gated (by voltage) and forming an ion-selective pore [17]. Voltage‑gated channels are critical to normal neuromuscular transmission and disruption of their normal function can lead to rapid paralysis. Toxins that target these components [17], are a valuable tool for understanding the structure and function of ion channels [13]. In this review, we will only refer to K_V and Na_V channel toxins, from all the ion channel toxins known.

Besides neurotoxins, cytolytic sea anemone toxins attracted considerable interest starting in the 1970s. The first report was of a phospholipase A in Aiptasia pallida venom [39]. Equinatoxin (Eqt), named following its source, the sea anemone Actinia equina [40], was the first actinoporin described in cnidarians.

As referred previously, not all the toxins are related to nematocyst [10,11]. According to Anderluh and co-workers [41], those that are, have a common signal directing them to a maturing cnidocyst [41]. Between the signal peptide and mature region, toxins contain a propart of 9–17 residues long, always ending with Lys-Arg [41]. The propart is composed mainly of polar and negatively charged amino acids, having the role to conduct the toxin to the nematocyst. However, in more recent works it was showed that Nv1 from Nematostella vectensis [42] and Anthopleura elegantissima toxins have proparts that also end in a Lys-Arg tandem but are not localized in the nematocysts. This suggests that the propart may have another role other than conducting toxins to the nematocysts. In another work, it was found that this cleavage sequence is not always conserved. Indeed, in the AvTX-20 (belonging to Cytolysins Type II) from Actineria villosa the propart terminate with a Lys-Lys sequence [43].

When scientists first extracted venoms from cnidarians, they started with species that had the more widespread toxic effects known. Thus, medusas have been the first organism studied, and all the subsequent works have used the protocol of Bloom and co-workers in 1998 as the major technique [138]. In that work, jellyfish tentacles were removed, stored in seawater, and vigorously shaken daily and let to settle to allow the release of the nematocysts. To recover nematocysts, the solution was filtered through a fine sieve. Glass beads, sonication or even freeze-thaw cycles were used with subsequent centrifugation of the solution to remove the cell debris from the venom.

Regarding the venom extraction in sea anemones, many techniques are employed since then and the techniques for extracting the venoms were obviously improved. In general, several techniques allow the extraction of the sea anemones venoms, which can be removed from the entire animal body or just from parts of the body, such as tentacles, acontia or acrorhagi. The tissue can be processed immediately, frozen or freeze-dried. Moreover, the venom can be obtained without animal injury, just by electric stimulation or gently squeezing the sea anemones. The majority of protocols use water to extract the venom, nonetheless there are other solutions that can be used, such as acetone. Most of the works purify the venom after being obtainment, by gel chromatography, followed by reverse phase HPLC. Some of them even go for SDS-PAGE. This chapter does not represent an exhaustive explanation of all the techniques used and improvements since the 1970s but instead provides a brief overview of the most used techniques to make sea anemones venom extractions, so that beginners can have a compilation of them. Table 6, summarizes the most used protocols. It contains the species used in the paper referenced, the tissue type and amount that scientists used, or the number of individuals collected if they have not sacrificed the animal, the technique used for tissue storage, the solvent used for venom extraction, the mechanical treatment used for venom extraction, the technique used for toxin recovery and reference.

While there have not been any paper evaluating the technique’s merit, the procedures that do not injury the animal are better for the obvious reason plus the venom seem to be better in terms of purity and it can be achieved also in good amounts. Moreover, frozen the specimens at −20 °C is also practicable, and good results have been obtained even without using −80 °C freezers, so this conservation method would be a good starting point. Using water as a solvent and a blender, followed by centrifugation, is an easy technique not time consuming nor demanding in terms of materials or skills, thus can be also used as a starting point for venom extraction. Regarding the toxins extraction only from the nematocysts, this can be accomplished if subjecting these structures to sonication or freeze-thaw cycles to burst and release the content.

The first toxins to be studied at genomic level were Equinatoxins and they proved to be intronless. Similarly, Or-A and Or-G also do not contain introns [120]. Afterwards it was found that some toxins, namely Cytolysins Avt-I and Pstx-20A, have three exons (two introns). Moreover, they are coded by at least two genes. Such gene arrangement is not exclusive of Cytolysins. The neurotoxic Clx-I and II, and HmK also have genes that are interrupted by two introns and their exon-intron organization is quite similar to the Avt-I genome structure. The introns-exon junctions that are typical donor and acceptor splice sites have followed the GT/AG rule, in which the introns begin with GT and end with AG [131]. In the work of Gendeh and co-workers [142] on HmK, a similar organization on introns-exon junction in scorpion toxins has been reported, suggesting that molecules with similar functions have similar organization at genomic level, therefore implying a common evolutionary path.

More than five equinatoxins genes are found and two isoforms of Equinatoxin-I [114]. ATX-II is encoded by at least seven genes [20]. In 2008, Wang and co-workers [130] showed that magnificalysins (HMgs) are also encoded by a multigene family, with each member encoding an isoform. They cloned more than 50 genes, all intronless. From the Nematostella vectensis whole genome release, it was found 13 genes that encode for the Nv1 toxin. Thus, toxins that are encoded by gene families may be more common than previously believed. Indeed, alternative splicing is not commonly assumed for toxins that have not yet the gene(s) sequenced.

The eight genes that code for the same toxin, Nv1, are arranged more or less sequentially and this supports the concerted evolution theory [143]. This theory is corroborated by what it happens in yeast- unequal crossover. In this unusual phenomenon, the sequence of genes is homogenized through unequal crossing over and gene conversion, resulting in an arrangement where two gene family members from one species are more similar to one another, than to their corresponding homologues in other species [143]. The advantage of this mechanism, having several copies of the same gene, is to produce rapidly a huge amount of venom. The nematocyst is discarded after each discharge, and the absence of a specialized venom organ/gland provide emphasis to this hypothesis. They also add that a multigene family, give organisms two more advantages: (i) the rapid transmission of advantageous mutations and (ii) the prevention of the loss of a highly effective toxin [20]. In the same work, some putative toxins in Anemonia viridis and Actinia equina were found to evolve in the opposite manner, by accelerated evolution, similar to what happens commonly in other venomous animals. Some toxins may escape from the concerted evolution process, diverging rapidly and with mutations being influenced by its selective value and neutral genetic drift [20]. Diversifying selection or Darwinian selection promotes the fixation of non-synonymous substitutions and ‘‘accelerates’’ the diversification of related sequences. This high substitution rate is typical to the region encoding the mature toxin. In contrast, the regions encoding the signal peptide and propart, which are involved in secretion, are usually highly conserved. In fact, in another work, the analysis of ATX-I and ATX-III from Anemonia viridis, revealed that besides the differences between the two toxins and 3D structure, the signal was conserved. This has likely been generated by gene fusion and advantageous in transcript stability and intracellular trafficking and secretion [143].

More recently, the genes encoding Kunitz-type toxins from Heteractis crispa have been studied [144]. Kunitz-type proteins are encoded by four distinct gene families (GS-, RG-, GG-, and GN-gene families). In one family studied (GS), several homologues peptides were found. Moreover, the Open Reading Frame is interrupted by a single intron located at the middle of the signal peptide. The scientists suggest that the gene family in case evolved through gene tandem duplication flowed by adaptive divergence of the reactive site (a particularly group of amino acids). Furthermore, this evolution seem to be lineage-specific, increasing the ability of Heteractis crispa to interact with multiple preys and foes [144].

Genes and transcripts of toxins are determined by PCR and degenerate primers by RACE, usually from the cDNA sequence and cloning.

Descriptions of sea anemone protein structures involved in venom activity have been determined by nuclear magnetic resonance (NMR) or X-ray crystallography. With the exception of three Actinoporins (Equinatoxin-II, Sticholysin-II and Fragaceatoxin C), all the other toxins were determined by solution NMR. In the same manner, all the toxins have only one chain, except the Actinoporins that have two, or six in the case of fragaceatoxin. Table 7 shows all the sea anemone toxins with three-dimensional structures described, the species from which it was purified, the type, the resolution method, the number of chains of the molecule and the amino acid number, and the RCSB PDB ID.

The first 3D protein structures of sea anemone were obtained in the 80’s by NMR analysis, for the Anthopleurin-A and ATX-IA [145,146]. However, the first pore-forming toxin to be studied was the Equinatoxin-II in 2001.

Sea anemone Actinoporins, Equinatoxin-II and Sticholysin-II, display an extremely similar structural organisation. The molecule is composed of a tightly folded β-sandwich core flanked on two sides by α-helices (Figure 6). The first 30 amino acids encompass one of the helices. This is the only part of the molecule able to undergo a conformational change without any structural change of the β-sandwich. A prominent patch of aromatic amino acids is located on the bottom of the molecule. It comprises a completely exposed Trp¹¹² (in EqtII), which was shown to participate in the initial binding of the toxin to the lipid membrane [124]. Comparing Fragaceatoxin C (FraC) with these two other actinoporins, it has a β-sandwich core flanked by three helices and the N-terminal domain is more “detached” from the protein core [147,148].

As previously referred, Actinoporins form pores in membranes by rearranging themselves in a four monomers structure [149]. For this purpose, four regions seem to be important in this step: A cluster of aromatic residues, a phosphocholine binding site, an array of basic amino acids, and the N-terminal α-helix. Initial binding of the soluble monomers to the membrane is accomplished by the cluster of aromatic amino acids, the array of basic residues, and the phosphocholine binding site. Then, the N-terminal α-helix detaches from the β-sandwich, extends, and lies parallel to the membrane. Simultaneously, oligomerization occurs. Finally, the extended N-terminal α-helix penetrates the membrane to build a toroidal pore [150].

The Na_V channel toxin from Anemonia viridis has a complete different conformation. ATX III adopts a compact structure, being the smallest of the structures, containing four reverse turns (a distorted type I β-turn, a type I β-turn, and an inverse γ-turn) and two other chain reversals, but no regular α-helix or β-sheet (Figure 7). In this molecule, several of the residues most affected by aggregation are located on the surface of the molecule [151], which cluster on one hemisphere and include a patch of hydrophobic residues only partially exposed [45].

With the analysis of the BgK structure, Dauplais and co-workers [152] verified that toxins with different structures from different organisms, like BgK from a sea anemone and ChTX, charybdotoxin (2CRD-PDB Accession Number), from a scorpion, which bind the K_V channel, have conserved a dyad. Such a dyad is composed of an essential Lys assisted by a more or less distant aromatic residue, whose precise nature (Tyr or Phe) and location may differ from one toxin to another. This fact suggests a convergent functional evolution for these small proteins [152]. See Figure 8 for structural differences between BgK and ChTx toxins and Figure 9 for conserved dyad between the two toxins.

The structure of CgNa, which was solved by NMR spectroscopy, is somewhat atypical and display significant homology with both type I and type II anemone toxins in amino acid sequence. CgNa also displays a considerable number of exceptions to the canonical structural elements that are thought to be essential for the activity of this group of toxins. Furthermore, unique residues, as Asp³⁶, Glu³⁷ and Glu⁴³ in CgNa (instead of Trp⁴⁵ in Anthopleurin-B), define a characteristic structure with strong negatively charged surface patches (Figure 10). These patches disrupt a surface-exposed cluster of hydrophobic residues present in all anemone-derived toxins described to date. CgNa preferentially binds to TTX-S (tetrodotoxin-sensitive) Na_V channels in the resting state. The specific structural features of CgNa may explain its weaker inhibitory capacity when compared with the other type I and II anemone toxins [154].

ATX-IA, Anthopleurin-A and B and Sh1 are constituted by four-stranded β-sheets. In the ATX-IA they are connected by two loops and there is an additional flexible loop consisting of 11 residues [146]. In Anthopleurin-A and Sh1 they are connected by three loops [155,156] and in Anthopleurin-B by several β-turns [157]. Anthopleurin-A, B and Sh1 have antiparallel β-sheets. APETx1 and BDS-I have three-stranded anti-parallel β-sheets. In addition, BDS-I has one more mini antiparallel β-sheet at the N-terminus. The β-sheet is connected by a long exposed loop [158].

The calculated structure of APETx1 belongs to the disulfide-rich all-β structural family, in which a three-stranded anti-parallel β-sheet is the only secondary structure. APETx1 is the first Ether-a-go-go effector discovered to fold in this way [159]. The hERG (the human Ether-à-go-go-Related Gene) is a gene (KCNH2) that codes for a protein known as K_V11.1 potassium ion channel. This ion channel (sometimes simply denoted as “hERG”) is best known for its contribution to the electrical activity of the heart that coordinates the heart’s beating.

The structures of the K_V channel toxins, as BDS-I, APETx1 and APETx2 are similar to those of the Na_V channel toxins such as Anthopleurin-A, but quite different from the ShK/BgK family of K_V channel toxins. This evidence clearly shows that sea anemones are capable of using a common structural scaffold to create blockers of distinct targets, e.g. Anthopleurin-A, APETx1 and APETx2 act on Na_V channel, hERG and ASIC channels, respectively, while also using different scaffolds (all-β in APETx1 vs. all-α in ShK) to block similar channels (hERG and K_V1, respectively) [19].

In spite of the recent increasing effort to study cnidarians venoms, much more is yet to be done to characterize these compounds in this diverse group of animals. The venom from each species of cnidarians is supposed to contain around 100 compounds, but not more that 1% is currently known even in the better studied species. Indeed, a recent work described 156 peptide venom compounds in a single species [35]. In this sense, toxins such as Actinoporins or PLA2s could be particularly interesting. First, they are less studied than other toxins in cnidarians and second, concerning PLA2s, they are a wide group of toxins also encountered in other animals, as in the better studied snakes. However, much more effort is also needed to pursue the study of ion channel toxins, which will allow a better understanding, not only of the diversity of those toxins, but also of the function of ion channels.

Another field starting to gain relevance is toxin gene detection. The increase in availability of genome sequences for venomous animals or cnidarians in general, added to the development of deep sequencing technology, will enable in depth study of genes encoding toxins. Such genomic studies will not only shed light on the evolutionary mechanisms influencing venom evolution but also, more broadly, on the genetic processes that underline the evolution of novel functionalities.

Cnidarians have impressive strategies for locomotion, feeding and reproduction. Its detailed study may allow unraveling the key for new medical drugs, as well as better understand the diversification of the molecular novelties of these unique metazoan species. For all such reasons, the study of cnidarians in whatever field is of great importance. From genetic and ecological studies to more applied pharmacological and toxicological assessments, these soft body animals should be a target of future scientific research.