Animal eyes have photoreceptor cells that contain light-sensitive molecules called opsins. Although all animal photoreceptor cells of this kind share a common origin, the cells found in different organisms can differ considerably. The photoreceptor cells in flies, squids and other invertebrates store a type of opsin called r-opsin in thin projections on the surface known as microvilli. On the other hand, the visual photoreceptor cells in human and other vertebrate eyes transport another type of opsin (known as c-opsin) into more prominent extensions called cilia.

It has been suggested that the fly and vertebrate photoreceptor cells represent clearly distinct evolutionary lineages of cells, which diverged early in animal evolution. However, several organisms that are more closely related to flies than to vertebrates have eye photoreceptor cells with cilia. Do all eye photoreceptors with cilia have a common origin in evolution or did they emerge independently in vertebrates and certain invertebrates?

The photoreceptor cells of a marine mollusc called Leptochiton asellus, are unusual because they bear both microvilli and cilia, suggesting they have intermediate characteristics between the two well-known types of photoreceptor cells. Previous studies have shown that these photoreceptor cells use r-opsin, but Vöcking et al. have now detected the presence of an additional opsin in the cells. This opsin is a member of the recently discovered xenopsin family of molecules. Further analyses support the findings of previous studies that suggested this type of opsin emerged early on in animal evolution, independently from c-opsin. Other invertebrates that have cilia on their eye photoreceptors also use xenopsin and not c-opsin.

The findings of Vöcking et al. suggest that, in addition to c-opsin and r-opsin, xenopsin has also driven the evolution of photoreceptor cells in animals. Eye photoreceptor cells in invertebrates with cilia probably share a common origin with the microvilli photoreceptor cells that is distinct from that of vertebrate visual cells. The observation that two very different types of opsin can be produced within a single cell suggests that the molecular processes that respond to light in photoreceptor cells may be much more complex than previously anticipated. Further work on these processes may help us to understand how animal eyes work and how they are affected by disease.

Animal eyes are amongst the best-investigated sensory organs and are subject to studies not only in sensory biology, but also in general molecular, developmental and evolutionary biology. Comparative investigations yield insights into the diversity of light-detection mechanisms and their function across animals. Further, the integrative data sets on eyes and their cellular components has had considerable impact on the characterization of different kinds of light-detecting cells, and on concepts how to trace their evolution and that of the organs they constitute (Arendt, 2003; Arendt et al., 2016; Oakley and Speiser, 2015; Wagner, 2014). These studies have dealt with the general question of whether gradual change or rather integration and reshuffling of modules that also exist elsewhere is the main driving force in the evolution of light-detecting cell types.

Traditionally, photoreceptor cells (PRCs) have been characterized mainly by their general electrophysiological response and their fine structure, which reveal clear differences between the two main systems of interest, the eyes of insects and those of vertebrates. The PRCs in insects have been shown to enlarge the sensory area by microvilli and to depolarize in response to light, whereas vertebrate eyes are hyperpolarizing and bear a modified cilium (Eakin, 1963; 1968, 1979). These findings initiated a wealth of comparative studies and resulted in pro-longed debates on whether a few conserved lineages of animal eye PRCs exist or whether both those PRCs andwhole eyes evolved multiple times independently (Eakin, 1982; von Salvini-Plawen, 1982; Vanfleteren, 1982).

Subsequent data on molecular physiology and transcriptional regulation of cell specification allowed much more detailed characterization of PRCs,suggesting that two conserved PRC types are indeed the main sensory components of animal cerebral eyes: (a) microvillar depolarizing r-opsin-expressing PRCs, which signal via the Gq-mediated IP3 cascade, and (b) ciliary hyperpolarizing c-opsin-expressing cells, which signal via the Gi/t-mediated cGMP cascade (Arendt et al., 2002a, 2004; Fain et al., 2010; Fernald, 2006; Gehring, 2014). This evolution-based classification of PRCs became a sound basis for many kinds of comparative eye research and for ideas about how animal eyes evolved.

The r-opsin+ type of PRCs constitutes the main visual elements in many protostome cerebral eyes, but they are also present in the ganglion cell layer of vertebrate eyes, where they are involved in the pupillary reflex and the entrainment of the circadian clock (Hattar et al., 2003; Koyanagi et al., 2005; Panda et al., 2002). Ciliary c-opsin+ PRCs on the other hand are predominantly known as the visual PRCs of vertebrate eyes and the frontal eye of cephalochordates (Vopalensky et al., 2012), while their role in protostomes is controversially discussed. In protostomes, c-opsins were initially reported only in arthropod and annelid deep brain photoreceptors (Arendt et al., 2004; Velarde et al., 2005). It was assumed, therefore, that this visual pigment was ancestrally employed by unpigmented brain PRCs that were fulfilling non-visual functions and were only secondarily integrated into the eyes of chordates (Arendt et al., 2004; Vopalensky et al., 2012). Reports on the presence of ciliary PRCs and c-opsins in the eyes of protostomes challenged this view and also the concept of gradually evolving ancient PRC types in animal eyes. Accordingly, a rather flexible employment of different opsin types by eye photoreceptor cells or frequent gain and loss of photoreceptor cell types in cerebral eyes during the evolution of bilaterian animals were suggested (Passamaneck et al., 2011).

Advances in opsin phylogeny, however, suggest that several protostome sequences, which share characteristic sequence motifs with c-opsins and were initially classified as such, fall into their own group, now termed xenopsins (Ramirez et al., 2016). In this situation, it is highly desirable to obtain deeper insights into the evolutionary history of c-opsins and xenopsins and their employment in animal photoreceptor cells, especially in those of protostome invertebrates. By deeply mining public and new data, we find xenopsins to be present in several taxa of protostome invertebrates. Although we did not find even a single case of co-existence of c-opsins and xenopsins in any organism, our phylogenetic and gene structure analyses strongly suggest that xenopsins and c-opsins are two distinct opsin subgroups. A new xenopsin from Leptochiton asellus, a member of the basally branching mollusk group of chitons, is found to be co-expressed with r-opsin in PRCs bearing both microvilli and cilia. This raises interesting questions about the light transduction and molecular physiology of the cells and their evolutionary origin. Recompilation of existing and new data challenges the dichotomous distinction of bilaterian eye PRCs into ciliary c-opsin+ and microvillar r-opsin+ cell types, but nevertheless supports the view that conserved lineages of eye PRCs exist. We suggest that a highly plastic PRC type of mixed microvillar/ciliary organization and frequent lineage-specific loss of c-opsins and xenopsins were important players in animal eye evolution.

In vertebrates, c-opsins drive the important process of vision, as they constitute the visual pigments of the retinal rods and cones and they serve additional functions when expressed in the pineal or in deep brain PRCs. Thus, the identification of c-opsins in protostome invertebrates provided the possibility to explore the role of c-opsin+ PRCs in distant relatives and to uncover the evolutionary roots of vertebrate vision. For a long time, only very few protostome c-opsins were known and these unambiguously group within the clade of c-opsins. This narrowed down the choice of potential study subjects. Basically, there were few pteropsins from arthropods and one sequence from the annelid Platynereis dumerilii (Arendt et al., 2004; Eriksson et al., 2013; Velarde et al., 2005).

In addition, new protostome sequences, which resemble c-opsins in certain aspects, were not instantly included in broad opsin analyses and this led to weakly supported orthology assignments that suggested a broader distribution of c-opsins in protostome invertebrates. Seemingly, this is not the case. Initial evidence that the new sequences do not encode c-opsins, but are closer affiliated to cnidops, has been provided by Hering and Mayer (2014), though with low support. This new group included sequences of hitherto uncharacterized visual pigments from the mollusks Crassostrea gigas and Lottia gigantea, a sequence from the brachiopod T. transversa that originally was described as c-opsin (Passamaneck et al., 2011) , and a weakly associated sequence from Strongylocentrotus purpuratus (opsin2) that did not group with other major opsin sequence groups in other analyses (D'Aniello et al., 2015; Ooka et al., 2010; Raible et al., 2006). Probably because of the increased taxon sampling and formal tree reconciliation, the recent phylogenetic study of Ramirez et al. (2016) provides for the first time strong support for a new group of xenopsins, which are distinct from c-opsins. We show that irrespective of the remarkable mutually exclusive taxonomic distribution of xenopsins and c-opsins, this distinction does not rely on tree inference artifacts, as can be seen from the specific gene structures of both opsin types.

Our data reveal a broader occurrence of xenopsins in protostomes than hitherto anticipated, pointing towards a high relevance of this opsin type in animal physiology. Accordingly, xenopsins are present not only in mollusks, rotifers and brachiopods as already reported by Ramirez et al. (2016), but also in several flatworms and in Oweniidae, the first branch within annelids. The situation in nemerteans is not clear because the screened RNA-seq datasets are probably not representative due to the low total number of opsins found. In accordance with Ramirez et al. (2016), we have no evidence that xenopsins exist in deuterostomes. Strongylocentrotus purpuratus opsin2 is the only deuterostome sequence that has ever been suggested to show affinities to sequences now classified as xenopsins (Hering and Mayer, 2014). However, the high similarity of the gene structure to that of Lingula anatina peropsin A strongly supports a position of both sequences in the ancient small group of bathyopsins sensu Ramirez et al. (2016), which exclusively comprises only a few echinoderm and brachiopod sequences.

Nevertheless, the data provide clear evidence that xenopsins were already present in the last common ancestor of deuterostomes and protostomes. In the analysis of Ramirez et al. (2016) and in our study, xenopsins form the sister group to cnidops, and this may very likely reflect the evolutionary history. As the basal interrelationships of the major opsin groups were shown to be strongly affected by outgroup composition, taxon sampling, tree inference algorithms and parameters (compare Cronin and Porter, 2014; Feuda et al., 2012, 2014; Hering and Mayer, 2014; Porter et al., 2012), this hypothesis may, however, need further investigation. Unfortunately, gene structure data are not informative due to the lack of introns in cnidops. For this reason and in contrast to Ramirez et al. (2016), we apply the term xenopsin only to bilaterian sequences and not to cnidops. Nevertheless, the assumption that xenopsins emerged within protostomes would require a sister group relationship between xenopsins and another clade of protostome-specific opsins. This would need major rearrangements of opsin tree topology and is not supported by gene structure data. In the (unlikely) scenario that xenopsins do not turn out to be sister to cnidops, it remains to be determined whether xenopsins were already present in the ancestor of Bilateria or emerged in the last common ancestor of protostomes and deuterostomes. This depends on the final phylogenetic position of acoelomorphs in relation to Bilateria and on further upcoming sequence data. We could not detect any xenopsin or c-opsin in the screened RNA-seq data from seven acoelomorphs and one xenoturbellid.

According to our data, frequent lineage-specific loss of either xenopsins or c-opsins (or both) during animal evolution has to be assumed. To date, no explanation can be given as to why the two opsin groups do not co-occur in a single organism. The intriguing possibility remains that an underlying causal relation drove the evolution of a large protein family that way. This will be an interesting case to study on the molecular physiological and the genomic levels.

The observed co-expression of xenopsin and r-opsin in the eye PRCs of L. asellus is remarkable from a functional perspective. As a general observation, the vast majority of PRCs express only one type of visual opsin. In nearly all cases where more than one visual opsin was found in PRCs, the respective opsins belong to the same opsin subgroup and evoke the same phototransduction cascade. This has been shown for r-opsins in several arthropods, as for example in the eye PRCs of Limulus polyphemus (Katti et al., 2010), fiddler crabs (Rajkumar et al., 2010) and a butterfly (Arikawa et al., 2003), and for c-opsins in for example mice (Applebury et al., 2000), guinea pigs (Parry and Bowmaker, 2016), salamander (Isayama et al., 2014) and cichlid fishes (Dalton et al., 2015). In most cases, expansion or tuning of the visual spectrum is the suggested function. Co-expression of opsins belonging to different, evolutionary distinct opsin types is reported only very rarely, as in the case of the annelid P. dumerilii, where Go-opsins and r-opsin are employed in eye PRCs (Gühmann et al., 2015). Here, the main function likewise seems to be the expansion of the visual spectrum of the cells. Our findings in L. asellus point towards another direction. The L. asellus eye PRCs express all relevant elements of standard r-opsin-mediated GNAQ-dependent IP3 signaling (Vöcking et al., 2015). By contrast, the NKQ tripeptide motif present in L. asellus xenopsin is known to be involved in the GNAI- and cGMP-based signaling (Marin et al., 2000) of c-opsins and is similar to motifs in cnidops, some of which were shown to mediate GNAS- and cAMP-based signaling (Koyanagi et al., 2008; Liegertová et al., 2015). The eye PRCs might thus be able to integrate different light signals directly. To our knowledge, the only case known in which co-expressed opsins evoke competing signaling cascades is in the pineal of the lizard Uta stasburiana (Su et al., 2006). Functional analysis of the molecular physiology of the L. asellus eye PRCs may thus be of great interest to sensory biologists.

Xenopsins are seemingly able to enter cilia. This type of opsin is employed in the purely ciliary PRCs of the larval eyes of the brachiopod Terebratalia transversa (Passamaneck et al., 2011). We find it expressed in eye PRCs in L. asellus, whose microvilli employ r-opsin (Vöcking et al., 2015) but which also have cilia. The expression of Arf4, rab8, FIP3 and RPGR provides the possibility that these proteins are involved in xenopsin trafficking, as they are in c-opsin ciliary trafficking in vertebrate rods and cones (Liu et al., 1999; Wang and Deretic, 2014, 2015; Wang et al., 2012). As the C-terminal VxPx motif necessary for the Arf4-mediated transport of vertebrate c-opsins (Wang and Deretic, 2014) is missing in xenopsins, Arf4-mediated transport of xenopsin would require other protein interactions. Interestingly, the same holds true for ciliary transport of cnidops and of c-opsins from organisms other than vertebrates and tunicates, as the VxPx motif is only conserved in the latter opsin groups and not in cephalochordate, echinoderm or protostome c-opsins or cnidops, some of which have been shown to be expressed in ciliary PRCs (Arendt et al., 2004; Bielecki et al., 2014; Kozmik et al., 2008; Vopalensky et al., 2012). It may be that Arf4 binding to c-opsins other than those of vertebrates and tunicates, xenopsins and cnidops relies on other protein interactions as in the case of vertebrate c-opsins. As an alternative, Arf4-complex-mediated ciliary opsin trafficking has to be regarded as a chordate invention.

Our extensive data mining did not change the picture that arthropods and certain annelids are the only organisms in which insights can be obtained on the physiological role of the protostome counterparts of vertebrate visual pigments. C-opsins have been found in no other protostomes. As outlined above, xenopsin-employing PRCs are promising study subjects for many other questions in sensory biology. Furthermore, we propose that data on the employment of xenopsins have significant impact on the understanding of the evolution of animal eyes and of photoreceptor cell types in general. As discussed below, we regard the presence of xenopsins in ancestral eyes as a likely scenario. This implies that the occurrence of xenopsins in the eyes of protostomes is not due to the flexible recruitment of opsins in existing eyes during evolution or to frequent invasion of new kinds of photoreceptor cells into eyes, but rather to inherited employment in an ancestral eye photoreceptor cell type.

The wide distribution of r-opsin+ cells in the eyes of protostomes and deuterostomes, alongside the reported similarities in molecular physiology and the process of cell specification, have been interpreted by many studies as a sign of common evolutionary origin and the presence of r-opsin+ rhabdomeric PRCs in the eye of the last common ancestor of bilaterians (Arendt, 2008; Arendt et al., 2002a; Koyanagi et al., 2005; Lamb, 2013; del Pilar Gomez et al., 2009). Data on c-opsins, however, were interpreted in different ways. The pivotal role that c-opsins play in the rods and cones of vertebrate eyes is obvious and findings from the cephalochordate frontal eye suggest that c-opsin+ ciliary PRCs were the predominant eye sensory cells already in the last common ancestor of chordates (Vopalensky et al., 2012). In protostomes, c-opsins were initially described only in the brain (Arendt et al., 2004; Beckmann et al., 2015; Velarde et al., 2005), which corresponds to their broad occurrence in the brain of basally branching deuterostome c-opsins such as encephalopsins (Blackshaw and Snyder, 1999; Nissilä et al., 2012). Thus, it has been suggested that c-opsin+ PRCs did originally not form part of eyes, but were rather localized deeply in the brain and not associated with screening pigment (Arendt, 2008; Arendt et al., 2004; Lamb, 2013). They secondarily invaded the eyes of chordates, but stayed within the brain in protostomes.

The presence of ciliary PRCs revealed by structural studies of the eyes of some protostome groups (Purschke et al., 2006; Salvini-Plawen, 2008; Woollacott and Eakin, 1973; Woollacott and Zimmer, 1972) would contradict this scenario if those cells showedmolecular characteristics similar to those of c-opsin+ brain PRCs. Accordingly, the discovery of a c-opsin that is expressed in ciliary eye PRCs of a brachiopod put forward alternative explanations favoring a higher lability of PRC function and multiple independent recruitments of r-opsin+ and c-opsin+ PRCs for directional vision in animal eye evolution (Passamaneck et al., 2011). These data conflicts are resolved by the fact that all c-opsins that were previously reported from the visual cells of protostome eyes turned out to be xenopsins instead of c-opsins, according to Ramirez et al. (2016) and our study . The only exception relies on a PCR-based report of a true c-opsin in an onychophoran eye (Eriksson et al., 2013), but this is contradicted by a cellular expression analysis by Beckmann et al. (2015) showing that c-opsin is absent from the eye retina layer and only present deeply within the brain and the optical ganglia.

By contrast, all existing data on the expression of xenopsins point towards their presence in eyes. The opsin reported from brachiopod larval eyes (Passamaneck et al., 2011) is a xenopsin. We detected xenopsin in eye PRCs of a mollusk and in a few extraocular PRCs, which are regarded as derivatives of cerebral eyes (Vöcking et al., 2015). RNA-seq data from cephalopods likewise suggest the presence of xenopsins in eyes (Yoshida et al., 2015). One option to explain the appearance of xenopsin in protostome eyes is to assume repeated recruitment of xenopsins into this context. This implies the co-option of xenopsin by ancestral microvillar r-opsin+ eye PRCs in eyes like those of L. asellus. In purely ciliary eyes such as those in brachiopods, either the formerly r-opsin+ cells completely changed identity or the original PRCs were replaced by new ciliary xenopsin+ PRCs, or the whole eyes arose completely de novo while the ancestral microvillar eyes were reduced. Abrupt changes and new inventions would be important and frequent events in eye evolution.

Alternatively, assuming the presence of both r-opsin and xenopsin in ancestral eyes provides probably a simpler explanation (Figure 5). Notably, this is not in conflict with the absence of xenopsins in the eyes of well-studied protostomes such as arthropods, or certain annelids, as secondary loss of xenopsin in the respective lineages is clearly evident from sequence resources and opsin phylogeny. R-opsin and xenopsin may initially have been employed in different cells, but a very attractive hypothesis is that cellular co-expression of r-opsin and xenopsin, like that which we observed in the eyes of L. asellus, is not an exceptional case but is inherited from ancestral eye PRCs that employed both opsins and exhibited microvilli and cilia (Figure 5). The cerebral eye PRCs of mixed microvillar/ciliary organization described in some organisms (Blumer, 1996; Hughes, 1970; Zhukov et al., 2006) may have barely maintained the ancestral organization, whereas microvillar eye PRCs may have emerged by reducing the cilia in taxa in which xenopsin was secondarily lost and the only visual pigment that survived is that associated with microvilli. Vice versa, purely ciliary eye PRCs, as have been described in several protostomes (Blumer, 1994, 1999; Passamaneck et al., 2011; Woollacott and Eakin, 1973; Zimmer and Woollacott, 1993), may have formed coincidently with downregulation or complete loss of r-opsins. The ancestral r-opsin+ and xenopsin+ PRC type may have arisen within the protostome branch leading to lophotrochozoan organisms (variant A in Figure 5), but even a much earlier origin is conceivable (variant B in Figure 5) as the absence of xenopsins in deuterostome eyes can easily be explained by the obvious secondary loss of this visual pigment in these animals.

Figure 5

Download asset Open asset

The scenario outlined above links back to older debates about PRC evolution and the ancestral function and significance of vestigial cilia found in many invertebrate microvillar eye PRCs. The vestigial ciliary structures described vary in the length and in the structural organization of the microtubular axonem and the ciliary rootlet, and often only basal bodies are found (Eakin and Westfall, 1964; Eakin, 1972; Turbeville, 1991; Verger-Bocquet, 1992). These structures were often interpreted as remnants of the motile cilia of the epidermal cells from which the PRCs initially originated (Arendt et al., 2009; Eakin 1979, 1982; Mayer, 2006). Derivation from sensory cilia was only assumed in the context of a hypothesis proposing that animal eye PRCs originated multiple times within the different animal groups from inconspicuous cells enrolled in a diffuse dermal light sense (von Salvini-Plawen and Mayr, 1977; Salvini-Plawen, 2008). This thesis was contradicted by the upcoming molecular physiological and developmental data favoring conserved evolutionary lineages of r-opsin+microvillar and c-opsin+ ciliary PRC types (Arendt, 2008; Arendt et al., 2004; Fernald, 2006; Lamb, 2013; Shubin et al., 2009). However, the observed cellular co-expression of r-opsin and xenopsin in cells bearing microvilli and cilia now suggests that vestigial cilia in microvillar PRCs may indeed also go back to cilia with sensory functions. It is worth noting that the often inconspicuous primary cilia known from most kinds of vertebrate cells are considered to be sensory organelles that are important for proper cell function, intercellular signalling and development (Malicki and Johnson, 2017; Singla, 2006; Walz, 2017). Last but not least, the distinction between motile and sensory cilia is no longer regarded as clear cut. More and more motile cilia that also display sensory capabilities are being reported (Kleene and Van Houten, 2014; Shah et al., 2009; Warner et al., 2014) and the distribution across eukaryotes of molecular components relevant for locomotion and sensation suggests that cilia were enrolled in both functions upon their emergence (Bloodgood, 2010; Johnson and Leroux, 2010; Mitchell, 2007, 2017; Quarmby and Leroux, 2010). The same is conceivable for cilia in ancient photoreceptor cells.

As current data only show use of xenopsin, but not of c-opsin, in the ciliary structures of protostome cerebral eyes, the described scenario is in line with the hypothesis that the cerebral eyes of animals employ only few different PRC types. This scenario does not require the assumption of multiple recruitments of sensory cilia and/or visual pigments or PRCs by cerebral eyes or of multiple origins of those entities. Instead, the PRCs employed in prostostome cerebral eyes are rendered as representatives of a highly plastic cell type. The ratios of microvilli/cilia and of r-opsin/xenopsin, and in consequence the downstream targets and the physiological properties of the PRCs, may have changed repeatedly within evolutionary lineages unless either the r-opsin or the xenopsin genes were finally lost. Subsequent studies on xenopsin+ PRCs may thus strongly impact the understanding of the degree to which conserved cell types and the properties of sensory cells can change over time and of how these changes occur.

In conclusion, opsin phylogenetic analyses combined with gene-structure analysis strongly confirm the existence of an old clade of xenopsins, which is evolutionarily distinct from c-opsins. The physiological role of this opsin type is not yet understood and requires further studies, especially in the interesting case of co-expression with r-opsin. The taxonomic distribution and the available epression data suggest that xenopsins may play a pivotal role in tracing PRC and eye evolution in bilaterian animals. The dichotomous classification of eye PRCs into ciliary c-opsin+ and rhabdomeric r-opsin+ cell types may no longer be appropriate, and the documented presence of xenopsin in eye PRCs may be due to common ancestry. The described evolutionary scenario provides simple explanations for the origin of eye PRCs exhibiting both microvilli and cilia and for how purely ciliary PRCs may have emerged in protostome eyes, and it rises intriguing questions of how much conserved cell types can change over time.

1. 1. Vöcking O
   2. Kourtesis I
   3. Tumu S
   4. Hausen H

   (2017) Leptochiton asellus xenopsin gDNA

   Publicly available at NCBI Nucleotide (accession no: MF133514).

   https://www.ncbi.nlm.nih.gov/nuccore/MF133514
2. 1. Vöcking O
   2. Kourtesis I
   3. Tumu S
   4. Hausen H

   (2017) Platynereis dumerilii ciliary-opsin gDNA

   Publicly available at NCBI Nucleotide (accession no: MF133515).

   https://www.ncbi.nlm.nih.gov/nuccore/MF133515
3. 1. Vöcking O
   2. Kourtesis I
   3. Tumu S
   4. Hausen H

   (2017) Owenia fusiformis xenopsin1 cds

   Publicly available at NCBI Nucleotide (accession no: MF133511).

   https://www.ncbi.nlm.nih.gov/nuccore/MF133511
4. 1. Vöcking O
   2. Kourtesis I
   3. Tumu S
   4. Hausen H

   (2017) Owenia fusiformis xenopsin2 cds

   Publicly available at NCBI Nucleotide (accession no: MF133512).

   https://www.ncbi.nlm.nih.gov/nuccore/MF133512
5. 1. Vöcking O
   2. Kourtesis I
   3. Tumu S
   4. Hausen H

   (2017) Owenia fusiformis xenopsin3 cds

   Publicly available at NCBI Nucleotide (accession no: MF133513).

   https://www.ncbi.nlm.nih.gov/nuccore/MF133513
6. 1. Vöcking O
   2. Kourtesis I
   3. Tumu S
   4. Hausen H

   (2017) Leptochiton asellus ADP ribosylation factor 4A cds

   Publicly available at NCBI Nucleotide (accession no: MF133503).

   https://www.ncbi.nlm.nih.gov/nuccore/MF133503
7. 1. Vöcking O
   2. Kourtesis I
   3. Tumu S
   4. Hausen H

   (2017) Leptochiton asellus ADP ribosylation factor 4B cds

   Publicly available at NCBI Nucleotide (accession no: MF133504).

   https://www.ncbi.nlm.nih.gov/nuccore/MF133504
8. 1. Vöcking O
   2. Kourtesis I
   3. Tumu S
   4. Hausen H

   (2017) Leptochiton asellus Rab11 family-interacting protein 3

   Publicly available at NCBI Nucleotide (accession no: MF133505).

   https://www.ncbi.nlm.nih.gov/nuccore/MF133505
9. 1. Vöcking O
   2. Kourtesis I
   3. Tumu S
   4. Hausen H

   (2017) Leptochiton asellus regulatory factor X cds

   Publicly available at NCBI Nucleotide (accession no: MF133506).

   https://www.ncbi.nlm.nih.gov/nuccore/MF133506
10. 1. Vöcking O
    2. Kourtesis I
    3. Tumu S
    4. Hausen H

    (2017) Leptochiton asellus Forkhead box J1 cds

    Publicly available at NCBI Nucleotide (accession no: MF133507).

    https://www.ncbi.nlm.nih.gov/nuccore/MF133507
11. 1. Vöcking O
    2. Kourtesis I
    3. Tumu S
    4. Hausen H

    (2017) Leptochiton asellus myosin VIIa cds

    Publicly available at NCBI Nucleotide (accession no: MF133508).

    https://www.ncbi.nlm.nih.gov/nuccore/MF133508
12. 1. Vöcking O
    2. Kourtesis I
    3. Tumu S
    4. Hausen H

    (2017) Leptochiton asellus retinitis pigmentosa GTPase regulator cds

    Publicly available at NCBI Nucleotide (accession no: MF133509).

    https://www.ncbi.nlm.nih.gov/nuccore/MF133509
13. 1. Vöcking O
    2. Kourtesis I
    3. Tumu S
    4. Hausen H

    (2017) Leptochiton asellus Ras-related protein Rab-8

    Publicly available at NCBI Nucleotide (accession no: MF133510).

    https://www.ncbi.nlm.nih.gov/nuccore/MF133510
14. 1. Vöcking O
    2. Kourtesis I
    3. Tumu S
    4. Hausen H

    (2017) Echinoplana celerrima xenopsin

    Publicly available at NCBI Nucleotide (accession no: BK010203).

    https://www.ncbi.nlm.nih.gov/nuccore/BK010203
15. 1. Vöcking O
    2. Kourtesis I
    3. Tumu S
    4. Hausen H

    (2017) Prostheceraeus vittatus xenopsin A

    Publicly available at NCBI Nucleotide (accession no: BK010204).

    https://www.ncbi.nlm.nih.gov/nuccore/BK010204
16. 1. Vöcking O
    2. Kourtesis I
    3. Tumu S
    4. Hausen H

    (2017) Stylochus ellipticus xenopsin A

    Publicly available at NCBI Nucleotide (accession no: BK010205).

    https://www.ncbi.nlm.nih.gov/nuccore/BK010205
17. 1. Vöcking O
    2. Kourtesis I
    3. Tumu S
    4. Hausen H

    (2017) Prosthiostomum siphunculus xenopsin

    Publicly available at NCBI Nucleotide (accession no: BK010206).

    https://www.ncbi.nlm.nih.gov/nuccore/BK010206
18. 1. Vöcking O
    2. Kourtesis I
    3. Tumu S
    4. Hausen H

    (2017) Prostheceraeus vittatus xenopsin B1

    Publicly available at NCBI Nucleotide (accession no: BK010207).

    https://www.ncbi.nlm.nih.gov/nuccore/BK010207
19. 1. Vöcking O
    2. Kourtesis I
    3. Tumu S
    4. Hausen H

    (2017) Prostheceraeus vittatus xenopsin B2

    Publicly available at NCBI Nucleotide (accession no: BK010208).

    https://www.ncbi.nlm.nih.gov/nuccore/BK010208
20. 1. Vöcking O
    2. Kourtesis I
    3. Tumu S
    4. Hausen H

    (2017) Stylochus ellipticus xenopsin BA

    Publicly available at NCBI Nucleotide (accession no: BK010209).

    https://www.ncbi.nlm.nih.gov/nuccore/BK010209
21. 1. Vöcking O
    2. Kourtesis I
    3. Tumu S
    4. Hausen H

    (2017) Lineus longissimus GO-opsin

    Publicly available at NCBI Nucleotide (accession no: BK010210).

    https://www.ncbi.nlm.nih.gov/nuccore/BK010210

1. 1. Andrade SC
   2. Montenegro H
   3. Strand M
   4. Schwartz ML
   5. Kajihara H
   6. Norenburg JL
   7. Turbeville JM
   8. Sundberg P
   9. Giribet G

   (2014) Baseodiscus unicolor transcriptome

   Publicly available at NCBI Sequence Read Archive (accession no: SRR1505175).

   https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR1505175
2. 1. Andrade S

   (2012) Cephalothrix hongkongiensis transcriptome

   Publicly available at NCBI Sequence Read Archive (accession no: SRR618505).

   https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR618505
3. 1. Andrade SC
   2. Montenegro H
   3. Strand M
   4. Schwartz ML
   5. Kajihara H
   6. Norenburg JL
   7. Turbeville JM
   8. Sundberg P
   9. Giribet G

   (2014) Hubrechtella ijimai transcriptome

   Publicly available at NCBI Sequence Read Archive (accession no: SRR1505100).

   https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR1505100
4. 1. Cannon JT
   2. Vellutini BC
   3. Smith J 3rd
   4. Ronquist F
   5. Jondelius U
   6. Hejnol A

   (2016) Transcriptome of the nemertean Lineus longissimus

   Publicly available at NCBI Sequence Read Archive (accession no: SRR2682192).

   https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR2682192
5. 1. Halanych KM
   2. Kocot KM

   (2014) Malacobdella grossa transcriptome

   Publicly available at NCBI Sequence Read Archive (accession no: SRR1611560).

   https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR1611560
6. 1. Andrade SC
   2. Montenegro H
   3. Strand M
   4. Schwartz ML
   5. Kajihara H
   6. Norenburg JL
   7. Turbeville JM
   8. Sundberg P
   9. Giribet G

   (2014) Nipponnemertes transcriptome

   Publicly available at NCBI Sequence Read Archive (accession no: SRR1508368).

   https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR1508368
7. 1. Kocot K

   (2014) Paranemertes peregrina transcriptome

   Publicly available at NCBI Sequence Read Archive (accession no: SRR1611562).

   https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR1611562
8. 1. Whelan N

   (2014) Transcriptome sequencing of Tubulanus polymorphus

   Publicly available at NCBI Sequence Read Archive (accession no: SRR1611583).

   https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR1611583
9. 1. Egger B

   (2015) Echinoplana celerrima

   Publicly available at NCBI Sequence Read Archive (accession no: SRR1796488).

   https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR1796488
10. 1. Laumer C

    (2015) Geocentrophora applanata transcriptome

    Publicly available at NCBI Sequence Read Archive (accession no: SRR1955490).

    https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR1955490
11. 1. Egger B

    (2015) Itaspiella helgolandica

    Publicly available at NCBI Sequence Read Archive (accession no: SRR1797713).

    https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR1797713
12. 1. Laumer C

    (2015) Kronborgia cf. amphipodicola transcriptome

    Publicly available at NCBI Sequence Read Archive (accession no: SRR1976457).

    https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR1976457
13. 1. Egger B

    (2015) Microdalyellia schmidti

    Publicly available at NCBI Sequence Read Archive (accession no: SRR1797778).

    https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR1797778
14. 1. Laumer C

    (2015) Prorhynchus sp. I transcriptome

    Publicly available at NCBI Sequence Read Archive (accession no: SRR1980634).

    https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR1980634
15. 1. Laumer C

    (2015) Prostheceraeus vittatus transcriptome

    Publicly available at NCBI Sequence Read Archive (accession no: SRR2000268).

    https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR2000268
16. 1. Egger B

    (2015) Prosthiostomum siphunculus

    Publicly available at NCBI Sequence Read Archive (accession no: SRR1797833).

    https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR1797833
17. 1. Egger B

    (2015) Stenostomum sthenum

    Publicly available at NCBI Sequence Read Archive (accession no: SRR1801788).

    https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR1801788
18. 1. Laumer C

    (2015) Stylochus ellipticus larvae transcriptome

    Publicly available at NCBI Sequence Read Archive (accession no: SRR1980690).

    https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR1980690
19. 1. Cannon JT
    2. Vellutini BC
    3. Smith J 3rd
    4. Ronquist F
    5. Jondelius U
    6. Hejnol A

    (2016) Transcriptome of the nemertodermatid Ascoparia sp.

    Publicly available at NCBI Sequence Read Archive (accession no: SRR2682154).

    https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR2682154
20. 1. Cannon JT
    2. Vellutini BC
    3. Smith J 3rd
    4. Ronquist F
    5. Jondelius U
    6. Hejnol A

    (2016) Transcriptome of the acoel Diopisthoporus longitubus

    Publicly available at NCBI Sequence Read Archive (accession no: SRR3105704).

    https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR3105704
21. 1. Cannon JT
    2. Vellutini BC
    3. Smith J 3rd
    4. Ronquist F
    5. Jondelius U
    6. Hejnol A

    (2016) Transcriptome of the acoel Eumecynostomum macrobursalium

    Publicly available at NCBI Sequence Read Archive (accession no: SRR3105705).

    https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR3105705
22. 1. Cannon JT
    2. Vellutini BC
    3. Smith J 3rd
    4. Ronquist F
    5. Jondelius U
    6. Hejnol A

    (2016) Transcriptome of the acoel Isodiametra pulchra

    Publicly available at NCBI Sequence Read Archive (accession no: SRR2681926).

    https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR2681926
23. 1. Cannon JT
    2. Vellutini BC
    3. Smith J 3rd
    4. Ronquist F
    5. Jondelius U
    6. Hejnol A

    (2016) Transcriptome of the nemertodermatid Meara stichopi

    Publicly available at NCBI Sequence Read Archive (accession no: SRR2681155).

    https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR2681155
24. 1. Cannon JT
    2. Vellutini BC
    3. Smith J 3rd
    4. Ronquist F
    5. Jondelius U
    6. Hejnol A

    (2016) Transcriptome of the nemertodermatid Nemertoderma westbladi

    Publicly available at NCBI Sequence Read Archive (accession no: SRR2682004).

    https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR2682004
25. 1. Cannon JT
    2. Vellutini BC
    3. Smith J 3rd
    4. Ronquist F
    5. Jondelius U
    6. Hejnol A

    (2016) Transcriptome of the nemertodermatid Sterreria sp.

    Publicly available at NCBI Sequence Read Archive (accession no: SRR2682099).

    https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR2682099
26. 1. Cannon JT
    2. Vellutini BC
    3. Smith J 3rd
    4. Ronquist F
    5. Jondelius U
    6. Hejnol A

    (2016) Transcriptome of Xenoturbella bocki

    Publicly available at NCBI Sequence Read Archive (accession no: SRR2681987).

    https://trace.ncbi.nlm.nih.gov/Traces/sra/?run=SRR2681987

1. 1. Alten L
   2. Schuster-Gossler K
   3. Beckers A
   4. Groos S
   5. Ulmer B
   6. Hegermann J
   7. Ochs M
   8. Gossler A

   (2012) Differential regulation of node formation, nodal ciliogenesis and cilia positioning by Noto and Foxj1

   Development 139:1276–1284.

   https://doi.org/10.1242/dev.072728

   • Google Scholar
2. 1. Applebury ML
   2. Antoch MP
   3. Baxter LC
   4. Chun LL
   5. Falk JD
   6. Farhangfar F
   7. Kage K
   8. Krzystolik MG
   9. Lyass LA
   10. Robbins JT

   (2000)

   The murine cone photoreceptor: a single cone type expresses both S and M opsins with retinal spatial patterning

   Neuron 27:513–523.

   • PubMed
   • Google Scholar
3. 1. Arendt D
   2. Hausen H
   3. Purschke G

   (2009) The 'division of labour' model of eye evolution

   Philosophical Transactions of the Royal Society B: Biological Sciences 364:2809–2817.

   https://doi.org/10.1098/rstb.2009.0104

   • Google Scholar
4. 1. Arendt D
   2. Musser JM
   3. Baker CVH
   4. Bergman A
   5. Cepko C
   6. Erwin DH
   7. Pavlicev M
   8. Schlosser G
   9. Widder S
   10. Laubichler MD
   11. Wagner GP

   (2016) The origin and evolution of cell types

   Nature Reviews Genetics 17:744–757.

   https://doi.org/10.1038/nrg.2016.127

   • Google Scholar
5. 1. Arendt D
   2. Tessmar K
   3. de Campos-Baptista MI
   4. Dorresteijn A
   5. Wittbrodt J

   (2002a)

   Development of pigment-cup eyes in the polychaete Platynereis dumerilii and evolutionary conservation of larval eyes in Bilateria

   Development 129:1143–1154.

   • PubMed
   • Google Scholar
6. 1. Arendt D
   2. Tessmar K
   3. de Campos-Baptista MI
   4. Dorresteijn A
   5. Wittbrodt J

   (2002b)

   Development of pigment-cup eyes in the polychaete platynereis dumerilii and evolutionary conservation of larval eyes in bilateria

   Development 129:1143–1154.

   • PubMed
   • Google Scholar
7. 1. Arendt D
   2. Tessmar-Raible K
   3. Snyman H
   4. Dorresteijn AW
   5. Wittbrodt J

   (2004) Ciliary photoreceptors with a vertebrate-type opsin in an invertebrate brain

   Science 306:869–871.

   https://doi.org/10.1126/science.1099955

   • Google Scholar
8. 1. Arendt D

   (2003)

   Evolution of eyes and photoreceptor cell types

   The International Journal of Developmental Biology 47:563–571.

   • PubMed
   • Google Scholar
9. 1. Arendt D

   (2008) The evolution of cell types in animals: emerging principles from molecular studies

   Nature Reviews Genetics 9:868–882.

   https://doi.org/10.1038/nrg2416

   • Google Scholar
10. 1. Arikawa K
    2. Mizuno S
    3. Kinoshita M
    4. Stavenga DG

    (2003)

    Coexpression of two visual pigments in a photoreceptor causes an abnormally broad spectral sensitivity in the eye of the butterfly Papilio xuthus

    The Journal of Neuroscience : The Official Journal of the Society for Neuroscience 23:4527–4532.

    • PubMed
    • Google Scholar
11. 1. Bartolomaeus T

    (1992)

    Ultrastructure of the photoreceptors in the larvae of Lepidochiton cinereus (Mollusca, Polyplacophora) and Lacuna divaricata (Mollusca, Gastropoda)

    Microfauna Marina 7:215–236.

    • Google Scholar
12. 1. Beckmann H
    2. Hering L
    3. Henze MJ
    4. Kelber A
    5. Stevenson PA
    6. Mayer G

    (2015) Spectral sensitivity in onychophora (velvet worms) revealed by electroretinograms, phototactic behaviour and opsin gene expression

    Journal of Experimental Biology 218:915–922.

    https://doi.org/10.1242/jeb.116780

    • Google Scholar
13. 1. Bielecki J
    2. Zaharoff AK
    3. Leung NY
    4. Garm A
    5. Oakley TH

    (2014) Ocular and extraocular expression of opsins in the rhopalium of tripedalia cystophora (Cnidaria: Cubozoa)

    PLoS One 9:e98870.

    https://doi.org/10.1371/journal.pone.0098870

    • Google Scholar
14. 1. Blackshaw S
    2. Snyder SH

    (1999)

    Encephalopsin: A novel mammalian extraretinal opsin discretely localized in the brain

    The Journal of Neuroscience : The Official Journal of the Society for Neuroscience 19:3681–3690.

    • PubMed
    • Google Scholar
15. 1. Bloodgood RA

    (2010) Sensory reception is an attribute of both primary cilia and motile cilia

    Journal of Cell Science 123:505–509.

    https://doi.org/10.1242/jcs.066308

    • Google Scholar
16. 1. Blumer M

    (1994) The ultrastructure of the eyes in the veliger-larvae of Aporrhais sp. and Bittium reticulatum (mollusca, caenogastropoda)

    Zoomorphology 114:149–159.

    https://doi.org/10.1007/BF00403262

    • Google Scholar
17. 1. Blumer MJF

    (1996) Alterations of the eyes during ontogenesis inAporrhais pespelecani (mollusca, caenogastropoda)

    Zoomorphology 116:123–131.

    https://doi.org/10.1007/BF02526944

    • Google Scholar
18. 1. Blumer MJF

    (1999) Development of a unique eye: photoreceptors of the pelagic predator Atlanta peroni (gastropoda, heteropoda)

    Zoomorphology 119:81–91.

    https://doi.org/10.1007/s004350050083

    • Google Scholar
19. 1. Cannon JT
    2. Vellutini BC
    3. Smith J
    4. Ronquist F
    5. Jondelius U
    6. Hejnol A

    (2016) Xenacoelomorpha is the sister group to nephrozoa

    Nature 530:89–93.

    https://doi.org/10.1038/nature16520

    • Google Scholar
20. 1. Cardona A
    2. Saalfeld S
    3. Schindelin J
    4. Arganda-Carreras I
    5. Preibisch S
    6. Longair M
    7. Tomancak P
    8. Hartenstein V
    9. Douglas RJ

    (2012) TrakEM2 software for neural circuit reconstruction

    PLoS One 7:e318011.

    https://doi.org/10.1371/journal.pone.0038011

    • Google Scholar
21. 1. Choksi SP
    2. Lauter G
    3. Swoboda P
    4. Roy S

    (2014) Switching on cilia: transcriptional networks regulating ciliogenesis

    Development 141:1427–1441.

    https://doi.org/10.1242/dev.074666

    • Google Scholar
22. Book

    1. Cronin TW
    2. Porter ML

    (2014)

    The Evolution of invertebrate photopigments and photoreceptors

    In: Hunt D. M, Hankins M. W, Collin S. P, Marshall N. J, editors. Evolution of Visual and Non-Visual Pigments. Boston, MA: Springer. pp. 105–135.

    • Google Scholar
23. 1. D'Aniello S
    2. Delroisse J
    3. Valero-Gracia A
    4. Lowe EK
    5. Byrne M
    6. Cannon JT
    7. Halanych KM
    8. Elphick MR
    9. Mallefet J
    10. Kaul-Strehlow S
    11. Lowe CJ
    12. Flammang P
    13. Ullrich-Lüter E
    14. Wanninger A
    15. Arnone MI

    (2015) Opsin evolution in the ambulacraria

    Marine Genomics 24:177–183.

    https://doi.org/10.1016/j.margen.2015.10.001

    • Google Scholar
24. 1. Dalton BE
    2. Lu J
    3. Leips J
    4. Cronin TW
    5. Carleton KL

    (2015) Variable light environments induce plastic spectral tuning by regional opsin coexpression in the African cichlid fish, Metriaclima zebra

    Molecular Ecology 24:4193–4204.

    https://doi.org/10.1111/mec.13312

    • Google Scholar
25. 1. del Pilar Gomez M
    2. Angueyra JM
    3. Nasi E

    (2009) Light-transduction in melanopsin-expressing photoreceptors of Amphioxus

    PNAS 106:9081–9086.

    https://doi.org/10.1073/pnas.0900708106

    • Google Scholar
26. 1. Dubruille R
    2. Laurençon A
    3. Vandaele C
    4. Shishido E
    5. Coulon-Bublex M
    6. Swoboda P
    7. Couble P
    8. Kernan M
    9. Durand B

    (2002) Drosophila regulatory factor X is necessary for ciliated sensory neuron differentiation

    Development 129:5487–5498.

    https://doi.org/10.1242/dev.00148

    • Google Scholar
27. 1. Eakin R

    (1972)

    Photochemisty of Vision

    625–684, Structure of invertebrate photoreceptors, Photochemisty of Vision, 7.

    • Google Scholar
28. 1. Eakin RM
    2. Westfall JA

    (1964) Further observations on the fine structure of some invertebrate eyes

    Zeitschrift fur Zellforschung und Mikroskopische Anatomie 62:310–332.

    https://doi.org/10.1007/BF00339283

    • Google Scholar
29. Book

    1. Eakin RM

    (1963)

    Lines of evolution of photoreceptors

    In: Tyler A, Mazia D, editors. General Physiology of Cell Specialization. New York: McGraw-Hill. pp. 393–425.

    • Google Scholar
30. 1. Eakin RM

    (1968)

    Evolution of photoreceptors

    Evolutionary Biology 2:194–242.

    • Google Scholar
31. 1. Eakin RM

    (1979) Evolutionary significance of photoreceptors: in retrospect

    American Zoologist 19:647–653.

    https://doi.org/10.1093/icb/19.2.647

    • Google Scholar
32. Book

    1. Eakin RM

    (1982)

    Continuity and diversity in photoreceptors

    In: Westfall J, editors. Visual Cells in Evolution. New York: Raven Press. pp. 91–106.

    • Google Scholar
33. 1. Eriksson BJ
    2. Fredman D
    3. Steiner G
    4. Schmid A

    (2013) Characterisation and localisation of the opsin protein repertoire in the brain and retinas of a spider and an onychophoran

    BMC Evolutionary Biology 13:186.

    https://doi.org/10.1186/1471-2148-13-186

    • Google Scholar
34. 1. Fain GL
    2. Hardie R
    3. Laughlin SB

    (2010) Phototransduction and the evolution of photoreceptors

    Current Biology 20:R114–R124.

    https://doi.org/10.1016/j.cub.2009.12.006

    • Google Scholar
35. 1. Fernald RD

    (2006) Casting a genetic light on the evolution of eyes

    Science 313:1914–1918.

    https://doi.org/10.1126/science.1127889

    • Google Scholar
36. 1. Feuda R
    2. Hamilton SC
    3. McInerney JO
    4. Pisani D

    (2012) Metazoan opsin evolution reveals a simple route to animal vision

    PNAS 109:18868–18872.

    https://doi.org/10.1073/pnas.1204609109

    • Google Scholar
37. 1. Feuda R
    2. Rota-Stabelli O
    3. Oakley TH
    4. Pisani D

    (2014) The comb jelly opsins and the origins of animal phototransduction

    Genome Biology and Evolution 6:1964–1971.

    https://doi.org/10.1093/gbe/evu154

    • Google Scholar
38. 1. Fischer FP

    (1980)

    Fine structure of the larval eye of lepidochitona cinerea L

    Spixiana 3:53–57.

    • Google Scholar
39. 1. Gehring WJ

    (2014) The evolution of vision

    Wiley Interdisciplinary Reviews: Developmental Biology 3:1–40.

    https://doi.org/10.1002/wdev.96

    • Google Scholar
40. 1. Gühmann M
    2. Jia H
    3. Randel N
    4. Verasztó C
    5. Bezares-Calderón LA
    6. Michiels NK
    7. Yokoyama S
    8. Jékely G

    (2015) Spectral tuning of phototaxis by a go-opsin in the rhabdomeric eyes of platynereis

    Current Biology 25:2265–2271.

    https://doi.org/10.1016/j.cub.2015.07.017

    • Google Scholar
41. 1. Hammesfahr B
    2. Odronitz F
    3. Mühlhausen S
    4. Waack S
    5. Kollmar M

    (2013) GenePainter: a fast tool for aligning gene structures of eukaryotic protein families, visualizing the alignments and mapping gene structures onto protein structures

    BMC Bioinformatics 14:77.

    https://doi.org/10.1186/1471-2105-14-77

    • Google Scholar
42. 1. Hatje K
    2. Keller O
    3. Hammesfahr B
    4. Pillmann H
    5. Waack S
    6. Kollmar M

    (2011) Cross-species protein sequence and gene structure prediction with fine-tuned Webscipio 2.0 and Scipio

    BMC Research Notes 4:265.

    https://doi.org/10.1186/1756-0500-4-265

    • Google Scholar
43. 1. Hattar S
    2. Lucas RJ
    3. Mrosovsky N
    4. Thompson S
    5. Douglas RH
    6. Hankins MW
    7. Lem J
    8. Biel M
    9. Hofmann F
    10. Foster RG
    11. Yau K-W

    (2003) Melanopsin and rod–cone photoreceptive systems account for all major accessory visual functions in mice

    Nature 424:75–81.

    https://doi.org/10.1038/nature01761

    • Google Scholar
44. 1. Hering L
    2. Mayer G

    (2014) Analysis of the opsin repertoire in the tardigrade hypsibius dujardini provides insights into the evolution of opsin genes in panarthropoda

    Genome Biology and Evolution 6:2380–2391.

    https://doi.org/10.1093/gbe/evu193

    • Google Scholar
45. 1. Hughes HPI

    (1970) A light and electron microscope study of some opisthobranch eyes

    Zeitschrift fur Zellforschung und Mikroskopische Anatomie 106:79–98.

    https://doi.org/10.1007/BF01027719

    • Google Scholar
46. 1. Isayama T
    2. Chen Y
    3. Kono M
    4. Fabre E
    5. Slavsky M
    6. DeGrip WJ
    7. Ma J-X
    8. Crouch RK
    9. Makino CL

    (2014) Coexpression of three opsins in cone photoreceptors of the salamander Ambystoma tigrinum

    Journal of Comparative Neurology 522:2249–2265.

    https://doi.org/10.1002/cne.23531

    • Google Scholar
47. 1. Johnson JL
    2. Leroux MR

    (2010) cAMP and cGMP signaling: sensory systems with prokaryotic roots adopted by eukaryotic cilia

    Trends in Cell Biology 20:435–444.

    https://doi.org/10.1016/j.tcb.2010.05.005

    • PubMed
    • Google Scholar
48. 1. Katti C
    2. Kempler K
    3. Porter ML
    4. Legg A
    5. Gonzalez R
    6. Garcia-Rivera E
    7. Dugger D
    8. Battelle B-A

    (2010) Opsin co-expression in Limulus photoreceptors: differential regulation by light and a circadian clock

    Journal of Experimental Biology 213:2589–2601.

    https://doi.org/10.1242/jeb.043869

    • Google Scholar
49. 1. Kleene SJ
    2. Van Houten JL

    (2014) Electrical signaling in motile and primary cilia

    BioScience 64:1092–1102.

    https://doi.org/10.1093/biosci/biu181

    • Google Scholar
50. 1. Koyanagi M
    2. Kubokawa K
    3. Tsukamoto H
    4. Shichida Y
    5. Terakita A

    (2005) Cephalochordate melanopsin: evolutionary linkage between invertebrate visual cells and vertebrate photosensitive retinal ganglion cells

    Current Biology 15:1065–1069.

    https://doi.org/10.1016/j.cub.2005.04.063

    • Google Scholar
51. 1. Koyanagi M
    2. Takano K
    3. Tsukamoto H
    4. Ohtsu K
    5. Tokunaga F
    6. Terakita A

    (2008) Jellyfish vision starts with cAMP signaling mediated by opsin-Gs cascade

    PNAS 105:15576–15580.

    https://doi.org/10.1073/pnas.0806215105

    • Google Scholar
52. 1. Kozmik Z
    2. Ruzickova J
    3. Jonasova K
    4. Matsumoto Y
    5. Vopalensky P
    6. Kozmikova I
    7. Strnad H
    8. Kawamura S
    9. Piatigorsky J
    10. Paces V
    11. Vlcek C

    (2008) Assembly of the cnidarian camera-type eye from vertebrate-like components

    Proceedings of the National Academy of Sciences 105:8989–8993.

    https://doi.org/10.1073/pnas.0800388105

    • Google Scholar
53. 1. Kuwajima M
    2. Mendenhall JM
    3. Lindsey LF
    4. Harris KM

    (2013) Automated transmission-mode scanning electron microscopy (tsem) for large volume analysis at nanoscale resolution

    PLoS One 8:e59573–14.

    https://doi.org/10.1371/journal.pone.0059573

    • Google Scholar
54. 1. Lamb TD

    (2013) Evolution of phototransduction, vertebrate photoreceptors and retina

    Progress in Retinal and Eye Research 36:52–119.

    https://doi.org/10.1016/j.preteyeres.2013.06.001

    • Google Scholar
55. 1. Liegertová M
    2. Pergner J
    3. Kozmiková I
    4. Fabian P
    5. Pombinho AR
    6. Strnad H
    7. Pačes J
    8. Vlček Čestmír
    9. Bartůněk P
    10. Kozmik Z

    (2015) Cubozoan genome illuminates functional diversification of opsins and photoreceptor evolution

    Scientific Reports 5:11885.

    https://doi.org/10.1038/srep11885

    • Google Scholar
56. 1. Liu X
    2. Udovichenko IP
    3. Brown SD
    4. Steel KP
    5. Williams DS

    (1999)

    Myosin VIIa participates in opsin transport through the photoreceptor cilium

    The Journal of Neuroscience : The Official Journal of the Society for Neuroscience 19:6267–6274.

    • PubMed
    • Google Scholar
57. 1. Malicki JJ
    2. Johnson CA

    (2017) The cilium: cellular antenna and central processing unit

    Trends in Cell Biology 27:126–140.

    https://doi.org/10.1016/j.tcb.2016.08.002

    • Google Scholar
58. 1. Marin EP
    2. Krishna AG
    3. Zvyaga TA
    4. Isele J
    5. Siebert F
    6. Sakmar TP

    (2000) The amino terminus of the fourth cytoplasmic loop of rhodopsin modulates rhodopsin-transducin interaction

    Journal of Biological Chemistry 275:1930–1936.

    https://doi.org/10.1074/jbc.275.3.1930

    • Google Scholar
59. 1. Mayer G

    (2006) Structure and development of onychophoran eyes: what is the ancestral visual organ in arthropods?

    Arthropod Structure & Development 35:231–245.

    https://doi.org/10.1016/j.asd.2006.06.003

    • Google Scholar
60. Conference

    1. Miller MA
    2. Pfeiffer W
    3. Schwartz T

    (2010)

    Creating the CIPRES Science Gateway for inference of large phylogenetic trees

    Proceedings of the Gateway Computing Environments Workshop . pp. 1–8.

    • Google Scholar
61. Book

    1. Mitchell DR

    (2007)

    The evolution of eukaryotic cilia and flagella as motile and sensory organelles

    In: Jékely G, editors. Advances in Experimental Medicine and Biology. Springer. pp. 130–140.

    • Google Scholar
62. 1. Mitchell DR

    (2017) Evolution of cilia

    Cold Spring Harbor Perspectives in Biology 9:a028290.

    https://doi.org/10.1101/cshperspect.a028290

    • Google Scholar
63. 1. Nissilä J
    2. Mänttäri S
    3. Särkioja T
    4. Tuominen H
    5. Takala T
    6. Timonen M
    7. Saarela S

    (2012) Encephalopsin (OPN3) protein abundance in the adult mouse brain

    Journal of Comparative Physiology A 198:833–839.

    https://doi.org/10.1007/s00359-012-0754-x

    • Google Scholar
64. 1. Oakley TH
    2. Speiser DI

    (2015) How complexity originates: the evolution of animal eyes

    Annual Review of Ecology, Evolution, and Systematics 46:237–260.

    https://doi.org/10.1146/annurev-ecolsys-110512-135907

    • Google Scholar
65. 1. Oakley TH

    (2003) The eye as a replicating and diverging, modular developmental unit

    Trends in Ecology & Evolution 18:623–627.

    https://doi.org/10.1016/j.tree.2003.09.005

    • Google Scholar
66. 1. Ooka S
    2. Katow T
    3. Yaguchi S
    4. Yaguchi J
    5. Katow H

    (2010) Spatiotemporal expression pattern of an encephalopsin orthologue of the sea urchin Hemicentrotus pulcherrimus during early development, and its potential role in larval vertical migration

    Development, Growth & Differentiation 52:195–207.

    https://doi.org/10.1111/j.1440-169X.2009.01154.x

    • Google Scholar
67. 1. Panda S
    2. Sato TK
    3. Castrucci AM
    4. Rollag MD
    5. DeGrip WJ
    6. Hogenesch JB
    7. Provencio I
    8. Kay SA

    (2002) Melanopsin (Opn4) requirement for normal light-induced circadian phase shifting

    Science 298:2213–2216.

    https://doi.org/10.1126/science.1076848

    • Google Scholar
68. 1. Parry JWL
    2. Bowmaker JK

    (2016)

    Visual pigment coexpression in guinea pig cones:a microspectrophotometric study

    Investigative Ophthalmology & Visual Science 43:1662–1665.

    • Google Scholar
69. 1. Passamaneck YJ
    2. Furchheim N
    3. Hejnol A
    4. Martindale MQ
    5. Lüter C

    (2011) Ciliary photoreceptors in the cerebral eyes of a protostome larva

    EvoDevo 2:6.

    https://doi.org/10.1186/2041-9139-2-6

    • Google Scholar
70. 1. Plachetzki DC
    2. Degnan BM
    3. Oakley TH

    (2007) The origins of novel protein interactions during animal opsin evolution

    PLoS One 2:e1054.

    https://doi.org/10.1371/journal.pone.0001054

    • Google Scholar
71. 1. Porter ML
    2. Blasic JR
    3. Bok MJ
    4. Cameron EG
    5. Pringle T
    6. Cronin TW
    7. Robinson PR

    (2012) Shedding new light on opsin evolution

    Proceedings of the Royal Society B: Biological Sciences 279:3–14.

    https://doi.org/10.1098/rspb.2011.1819

    • Google Scholar
72. 1. Purschke G
    2. Arendt D
    3. Hausen H
    4. Müller MCM

    (2006) Photoreceptor cells and eyes in annelida

    Arthropod Structure & Development 35:211–230.

    https://doi.org/10.1016/j.asd.2006.07.005

    • Google Scholar
73. 1. Quarmby LM
    2. Leroux MR

    (2010) Sensorium: the original raison d'etre of the motile cilium?

    Journal of Molecular Cell Biology 2:65–67.

    https://doi.org/10.1093/jmcb/mjp036

    • Google Scholar
74. 1. Raible F
    2. Tessmar-Raible K
    3. Arboleda E
    4. Kaller T
    5. Bork P
    6. Arendt D
    7. Arnone MI

    (2006) Opsins and clusters of sensory G-protein-coupled receptors in the sea urchin genome

    Developmental Biology 300:461–475.

    https://doi.org/10.1016/j.ydbio.2006.08.070

    • Google Scholar
75. 1. Rajkumar P
    2. Rollmann SM
    3. Cook TA
    4. Layne JE

    (2010) Molecular evidence for color discrimination in the Atlantic sand fiddler crab, Uca pugilator

    Journal of Experimental Biology 213:4240–4248.

    https://doi.org/10.1242/jeb.051011

    • Google Scholar
76. 1. Ramirez MD
    2. Pairett AN
    3. Pankey MS
    4. Serb JM
    5. Speiser DI
    6. Swafford AJ
    7. Oakley TH

    (2016) The last common ancestor of most bilaterian animals possessed at least 9 opsins

    Genome Biology and Evolution 26:evw248.

    https://doi.org/10.1093/gbe/evw248

    • Google Scholar
77. 1. Rhode B

    (1992) Development and differentiation of the eye inPlatynereis dumerilii (Annelida, Polychaeta)

    Journal of Morphology 212:71–85.

    https://doi.org/10.1002/jmor.1052120108

    • Google Scholar
78. 1. Rosen M
    2. Stasek C
    3. Hermans C

    (1979)

    Ultrastructure and evolutionay significance of the ocelli in the larva of Katharina tunicata (Mollusca, Polyplacophora)

    The Veliger 22:173–178.

    • Google Scholar
79. 1. Saalfeld S
    2. Fetter R
    3. Cardona A
    4. Tomancak P

    (2012) Elastic volume reconstruction from series of ultra-thin microscopy sections

    Nature Methods 9:717–720.

    https://doi.org/10.1038/nmeth.2072

    • Google Scholar
80. 1. Salvini-Plawen L

    (2008)

    Photoreception and the polyphyletic evolution of photoreceptors (with special reference to mollusca)*

    American Malacological Bulletin 26:83–100.

    • Google Scholar
81. 1. Shah AS
    2. Ben-Shahar Y
    3. Moninger TO
    4. Kline JN
    5. Welsh MJ

    (2009) Motile cilia of human airway epithelia are chemosensory

    Science 325:1131–1134.

    https://doi.org/10.1126/science.1173869

    • Google Scholar
82. 1. Shubin N
    2. Tabin C
    3. Carroll S

    (2009) Deep homology and the origins of evolutionary novelty

    Nature 457:818–823.

    https://doi.org/10.1038/nature07891

    • Google Scholar
83. 1. Singla V

    (2006) The primary cilium as the cell's antenna: signaling at a sensory organelle

    Science 313:629–633.

    https://doi.org/10.1126/science.1124534

    • Google Scholar
84. 1. Song L
    2. Florea L

    (2015) Rcorrector: efficient and accurate error correction for Illumina RNA-seq reads

    GigaScience 4:48.

    https://doi.org/10.1186/s13742-015-0089-y

    • Google Scholar
85. 1. Su CY
    2. Luo DG
    3. Terakita A
    4. Shichida Y
    5. Liao HW
    6. Kazmi MA
    7. Sakmar TP
    8. Yau KW

    (2006) Parietal-eye phototransduction components and their potential evolutionary implications

    Science 311:1617–1621.

    https://doi.org/10.1126/science.1123802

    • PubMed
    • Google Scholar
86. Book

    1. Turbeville JM

    (1991)

    Nemertinea

    In: Bogitsh B. J, Harrison F. W, editors. Microscopic Anatomy of Invertebrates. New York: Wiley-Liss. pp. 285–328.

    • Google Scholar
87. Book

    1. Vanfleteren J

    (1982)

    A Monophyletic Line of Evolution? Ciliary Induced Photoreceptor Membrans

    In: Westfall J, editors. Visual Cells in Evolution. New York: Raven Press. pp. 107–136.

    • Google Scholar
88. 1. Velarde RA
    2. Sauer CD
    3. Walden KK
    4. Fahrbach SE
    5. Robertson HM

    (2005) Pteropsin: a vertebrate-like non-visual opsin expressed in the honey bee brain

    Insect Biochemistry and Molecular Biology 35:1367–1377.

    https://doi.org/10.1016/j.ibmb.2005.09.001

    • PubMed
    • Google Scholar
89. Book

    1. Verger-Bocquet M

    (1992)

    Polychaeta: Sensory Structures

    In: Gardiner S. L, Harrison F. W, editors. Microscopoic Anatomy of Invertebrates. New York: Wiley-Liss. pp. 181–196.

    • Google Scholar
90. Book

    1. von Salvini-Plawen L
    2. Mayr E

    (1977)

    On the evolution of photoreceptors and eyes

    In: Hecht M, Steere W, Wallace B, editors. Biology E. Volutionary. New York: Plenum Press. pp. 207–263.

    • Google Scholar
91. Book

    1. von Salvini-Plawen L

    (1982)

    On the polyphyletic origin of photoreceptors

    In: Westfall J, editors. Visual Cells in Evolution. New York: Raven Press. pp. 137–154.

    • Google Scholar
92. 1. Vopalensky P
    2. Pergner J
    3. Liegertova M
    4. Benito-Gutierrez E
    5. Arendt D
    6. Kozmik Z

    (2012) Molecular analysis of the amphioxus frontal eye unravels the evolutionary origin of the retina and pigment cells of the vertebrate eye

    PNAS 109:15383–15388.

    https://doi.org/10.1073/pnas.1207580109

    • Google Scholar
93. 1. Vöcking O
    2. Kourtesis I
    3. Hausen H

    (2015) Posterior eyespots in larval chitons have a molecular identity similar to anterior cerebral eyes in other bilaterians

    EvoDevo 6:1–14.

    https://doi.org/10.1186/s13227-015-0036-0

    • Google Scholar
94. Book

    1. Wagner GP

    (2014)

    Homology, Genes, and Evolutionary Innovation

    Princeton University Press.

    • Google Scholar
95. 1. Walz G

    (2017) Role of primary cilia in non-dividing and post-mitotic cells

    Cell and Tissue Research 369:11–25.

    https://doi.org/10.1007/s00441-017-2599-7

    • Google Scholar
96. 1. Wang J
    2. Deretic D

    (2014) Molecular complexes that direct rhodopsin transport to primary cilia

    Progress in Retinal and Eye Research 38:1–19.

    https://doi.org/10.1016/j.preteyeres.2013.08.004

    • Google Scholar
97. 1. Wang J
    2. Deretic D

    (2015) The Arf and Rab11 effector FIP3 acts synergistically with ASAP1 to direct Rabin8 in ciliary receptor targeting

    Journal of Cell Science 128:1375–1385.

    https://doi.org/10.1242/jcs.162925

    • Google Scholar
98. 1. Wang J
    2. Morita Y
    3. Mazelova J
    4. Deretic D

    (2012) The Arf GAP ASAP1 provides a platform to regulate Arf4- and Rab11-Rab8-mediated ciliary receptor targeting

    The EMBO Journal 31:4057–4071.

    https://doi.org/10.1038/emboj.2012.253

    • Google Scholar
99. 1. Warner JF
    2. McCarthy AM
    3. Morris RL
    4. McClay DR

    (2014) Hedgehog signaling requires motile cilia in the sea urchin

    Molecular Biology and Evolution 31:18–22.

    https://doi.org/10.1093/molbev/mst176

    • PubMed
    • Google Scholar
100. 1. Woollacott RM
     2. Eakin RM

     (1973) Ultrastructure of a potential photoreceptoral organ in the larva of an entoproct

     Journal of Ultrastructure Research 43:412–425.

     https://doi.org/10.1016/S0022-5320(73)90018-X

     • Google Scholar
101. 1. Woollacott RM
     2. Zimmer RL

     (1972) Fine structure of a potential photoreceptor organ in the larva of Bugula neritina (Bryozoa)

     Zeitschrift fur Zellforschung und Mikroskopische Anatomie 123:458–469.

     https://doi.org/10.1007/BF00335542

     • Google Scholar
102. 1. Yoshida MA
     2. Ogura A
     3. Ikeo K
     4. Shigeno S
     5. Moritaki T
     6. Winters GC
     7. Kohn AB
     8. Moroz LL

     (2015) Molecular Evidence for Convergence and Parallelism in Evolution of Complex Brains of Cephalopod Molluscs: Insights from Visual Systems

     Integrative and Comparative Biology 55:1070–1083.

     https://doi.org/10.1093/icb/icv049

     • PubMed
     • Google Scholar
103. 1. Yu X
     2. Ng CP
     3. Habacher H
     4. Roy S

     (2008) Foxj1 transcription factors are master regulators of the motile ciliogenic program

     Nature Genetics 40:1445–1453.

     https://doi.org/10.1038/ng.263

     • Google Scholar
104. 1. Zhukov VV
     2. Borissenko SL
     3. Zieger MV
     4. Vakoliuk IA
     5. Meyer-Rochow VB

     (2006) The eye of the freshwater prosobranch gastropod Viviparus viviparus: ultrastructure, electrophysiology and behaviour

     Acta Zoologica 87:13–24.

     https://doi.org/10.1111/j.1463-6395.2006.00216.x

     • Google Scholar
105. 1. Zimmer RL
     2. Woollacott RM

     (1993) Anatomy of the larva ofAmathia vidovici (Bryozoa: Ctenostomata) and phylogenetic significance of the vesiculariform larva

     Journal of Morphology 215:1–29.

     https://doi.org/10.1002/jmor.1052150102

     • Google Scholar

Author details

1. Oliver Vöcking

   1. Sars International Centre for Marine Molecular Biology, University of Bergen, Bergen, Norway
   2. Department of Ophthalmology, University of Pittsburgh, Pittsburgh, United States

   Contribution

   Data curation, Investigation, Visualization, Methodology, Writing—original draft, Writing—review and editing

   Competing interests

   No competing interests declared

2. Ioannis Kourtesis

   Sars International Centre for Marine Molecular Biology, University of Bergen, Bergen, Norway

   Contribution

   Investigation, Visualization, Methodology

   Competing interests

   No competing interests declared

3. Sharat Chandra Tumu

   Sars International Centre for Marine Molecular Biology, University of Bergen, Bergen, Norway

   Contribution

   Resources, Investigation, Methodology

   Competing interests

   No competing interests declared

4. Harald Hausen

   Sars International Centre for Marine Molecular Biology, University of Bergen, Bergen, Norway

   Contribution

   Conceptualization, Data curation, Software, Supervision, Validation, Investigation, Visualization, Methodology, Writing—original draft, Project administration, Writing—review and editing

   For correspondence

   harald.hausen@uib.no

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-2788-2835

• 3,590

  views

• 553

  downloads

• 56

  citations

Views, downloads and citations are aggregated across all versions of this paper published by eLife.