1. Evolutionary Biology
  2. Neuroscience
Download icon

The visual pigment xenopsin is widespread in protostome eyes and impacts the view on eye evolution

  1. Clemens Christoph Döring
  2. Suman Kumar
  3. Sharat Chandra Tumu
  4. Ioannis Kourtesis
  5. Harald Hausen  Is a corresponding author
  1. Sars International Centre for Marine Molecular Biology, University of Bergen, Norway
Research Article
  • Cited 1
  • Views 1,049
  • Annotations
Cite this article as: eLife 2020;9:e55193 doi: 10.7554/eLife.55193

Abstract

Photoreceptor cells in the eyes of Bilateria are often classified into microvillar cells with rhabdomeric opsin and ciliary cells with ciliary opsin, each type having specialized molecular components and physiology. First data on the recently discovered xenopsin point towards a more complex situation in protostomes. In this study, we provide clear evidence that xenopsin enters cilia in the eye of the larval bryozoan Tricellaria inopinata and triggers phototaxis. As reported from a mollusc, we find xenopsin coexpressed with rhabdomeric-opsin in eye photoreceptor cells bearing both microvilli and cilia in larva of the annelid Malacoceros fuliginosus. This is the first organism known to have both xenopsin and ciliary opsin, showing that these opsins are not necessarily mutually exclusive. Compiling existing data, we propose that xenopsin may play an important role in many protostome eyes and provides new insights into the function, evolution, and possible plasticity of animal eye photoreceptor cells.

Introduction

The photoreceptor cells (PRCs) in animal eyes are often classified according to their structure, that is depending on whether the sensory surface is enlarged by microvilli or by cilia (Eakin, 1979; Eakin, 1963; Eakin, 1968). The first type of PRCs in many protostomes was shown to depolarize in response to light and to employ rhabdomeric opsin (r-opsin) as a visual pigment, which signals via the Gαq mediated IP3 cascade opening TRP ion channels in the PRC membrane (Fain et al., 2010; Shichida and Matsuyama, 2009). In contrast, ciliary PRCs of vertebrate eyes are known to signal via the Gαi/t mediated cGMP cascade closing CNG channels and leading to a hyperpolarization. Since both are found in protostome and deuterostome animals and due to their distinct molecular signatures, it is assumed that these two kinds of PRCs were already present in the last common ancestor of bilaterian animals (Arendt, 2008; Arendt et al., 2004; Arendt et al., 2002; Gomez et al., 2009; Gehring, 2014; Nasi and Gomez, 2009; Panda et al., 2002). This classification of PRCs became a sound basis for comparative eye research from sensory biology to molecular physiological, developmental, and evolutionary biology. We present data suggesting that in protostomes, an additional second kind of ciliary PRCs is widespread and that this may be evolutionarily closer to microvillar PRCs than to vertebrate ciliary eye PRCs.

Recently, a new type of visual opsins, xenopsin, has been characterized. It shares important functional sequence motifs with ciliary opsins (c-opsins) and has been shown to signal most likely also via GαI in a flatworm (Rawlinson et al., 2019). Nonetheless, xenopsins and c-opsins do not group in phylogenetic analyses (Ramirez et al., 2016; Rawlinson et al., 2019; Vöcking et al., 2017) indicating a distinct evolutionary origin. Surprisingly, xenopsins and c-opsins are mutually exclusively distributed across the animal kingdom, which is difficult to explain from a genomic perspective and seemingly doubts the phylogenetic analyses. In this study, we report the first organism having both xenopsin and c-opsin. In congruence with thorough phylogenetic and gene structure analyses, this provides further support for a distinct evolutionary origin of these visual pigments.

Despite increasing knowledge on the presence of xenopsin in many animal groups, only very few data on cellular expression and function of xenopsin exist. So far it turned out to be this new opsin type and not c-opsin that is present in ciliary eye PRCs of larval brachiopods (Passamaneck et al., 2011; Vöcking et al., 2017) and in larval ciliary eye PRCs and adult extraocular ciliary PRCs in a flatworm (Rawlinson et al., 2019). Furthermore, xenopsin has been found coexpressed with r-opsin in eye PRCs exhibiting both microvilli and cilia in the larva of a mollusc (Vöcking et al., 2017), thereby raising the question, whether protostome eye PRCs had the potential to change between microvillar and ciliary organization during evolution.

To obtain a broader overview of the role of xenopsin in animal eyes, we investigated larva of the annelid Malacoceros fuliginosus (Claparède, 1868), and the bryozoan Tricellaria inopinata d'Hondt & Occhipinti Ambrogi, 1985 in which RNA-seq data pointed to the presence of xenopsin. We find it expressed in ciliary eye PRCs of the bryozoan larva, and we present unambiguous evidence that xenopsin enters the cilia and likely triggers the phototactic response of the larva. Further, we find xenopsin coexpressed with r-opsin in eye PRCs of the annelid larva similar to the earlier finding in a larval chiton (Vöcking et al., 2017). We propose that (1) Xenopsin is an important visual pigment in protostomes, (2) ciliary eye PRCs may not be of the same evolutionary origin in protostomes and deuterostomes, and (3) ciliary and microvillar eye PRCs may be evolutionarily linked in protostomes. The findings impact the current understanding of how animal eyes evolved and diversified and provides insights on the plasticity that cell types can exhibit in the course of evolution.

Results

Molecular phylogeny of animal xenopsins and c-opsins

We screened Tricellaria inopinata assembly one for opsins by blasting with a broad set of metazoan opsin sequences as query and successive reciprocal blast against Genbank. The sequences were further checked for the presence of the PFAM 7tm_1 domain and the residue Lys296, which is predictive for chromophore binding in opsins and for the NPXXY motif at positions 302–306 (Figure 1) contributing to signal transduction in G protein-coupled receptors. We blasted the hits against assembly two and elongated the sequences if longer hits were retrieved. We screened the transcriptomic resources of Malacoceros fuliginosus in the same manner, but only for the presence of xenopsins and c-opsins. We retrieved five hits from the T. inopinata assembly, which all gave xenopsins as first hits by reciprocal blast. Since we had evidence for contamination of the T. inopinata assembly (see Materials and methods), we cloned all sequences and tested them by ISH for expression in T. inopinata larva. Only one sequence gave positive signals and was further used in this study, while the others were no longer considered as they might be from other bryozoan species. Three sequences were retrieved from the M. fuliginosus assembly. After reciprocal blast against Genbank, one sequence gave c-opsins as first hits and the other two xenopsins. For further analyses, we kept the potential c-opsin and that potential xenopsin, for which we obtained positive results after in situ hybridization in larvae.

Figure 1 with 1 supplement see all
Conservation of functionally important motifs and residues in different opsin types.

Alignment of parts of the transmembrane domain VII and the cytosolic helix VIII of selected opsin sequences showing the conserved lysine 296 (K296) chromophore binding site and other conserved motifs important for opsin-G protein interaction like NPXXY and the tripeptide (NKQ in c-opsins and several xenopsins; HPK in r-opsins). The sequences investigated in this study are highlighted in blue.

We added the sequences and few recently described xenopsin sequences from the molluscs Sepia officinalis and Ambigolimax valentianus, the bryozoan Bugula neritina, the flatworm Maritigrella crozieri, and the chaetognath Pterosagitta draco to the opsin sequence set (https://doi.org/10.7554/eLife.23435.009) analyzed by Vöcking et al., 2017 and ran maximum likelihood and Bayesian phylogenetic analyses to study opsin molecular evolution with a focus on the relationships of xenopsins and c-opsins. All major opsin groups described by Ramirez et al., 2016, Vöcking et al., 2017, and Rawlinson et al., 2019 such as tetraopsins, r-opsins, cnidops, ctenopsins, c-opsins, and xenopsins were recovered with high support values (Figure 2, Figure 2—figure supplements 1 and 2). One sequence from M. fuliginosus falls into c-opsins, while another one falls into xenopsins. The opsin of T. inopinata likewise falls into xenopsin and groups with the sequence of the bryozoan Bugula neritina. The topology within the xenopsin clade suggests an early divergence of xenopsin in two clades xenopsin A and xenopsin B containing opsin from several animal groups similar as described by Vöcking et al., 2017 and Rawlinson et al., 2019. Yet, the support values for the two subclades are not as high as for xenopsisn as a whole and other large opsin groups. We tested robustness of the split into xenopsin A and B against changes in the outgroup by calculating trees of xenopsins only (unrooted) and trees with few cililary opsins, few cnidops and few c-opsins and cnidpos as outgroup. The split is retained in all cases with the exception of an outgroup composed of cnidops and c-opsins, where xenopsin B is a paraphyletic assemblage (Figure 2—figure supplements 58). The position of one brachiopod sequence (Lingula anatina melanopsin like XP 013397676.1) is not stable, in some cases it falls into xenopsin A, in others it groups with xenopsin B sequences or has a basal position. Accordingly, our data suggest an early diversification of xenopsins, but with moderate support only. Since M. fuliginosus xenopsin groups in all trees with xenopsin B representatives, we regard it as likely being the first known annelid xenopsin B. Several flatworm xenopsin B sequences stand out by strong modifications in the NPXXY and tripeptide motif (Figure 1—figure supplement 1) questioning the capability of the opsins to induce G-protein based light transduction. In difference, these motifs are conserved in the xenopsin of M. fuliginosus.

Figure 2 with 8 supplements see all
C-opsins and xenopsins display type-specific conserved gene structures.

Maximum Likelihood tree of opsin protein sequences (IQ-TREE, LG+F+R8). Labeled nodes have support values of SH-like approximate likelihood ratio test (blue dot) and ultrafast bootstrap ≥0.9 (purple dot), approximate Bayes test ≥0.98(yellow dot), and a posterior probability ≥0.95 (black dot) in a parallel Bayesian analysis (Phylobayes, DS-GTR + G, consensous of two out of three chains, 90,000 cycles). Intron positions (colored bars) are mapped on the un-curated protein sequence alignment, and introns conserved in position and phase are highlighted by bars spanning several sequences and labels for the intron position. The sequences investigated in this study are highlighted in blue. The xenopsins of M. fuliginosus and T. inopinata display xenopsin type gene structures. The c-opsin of M. fuliginosus groups with Platynereis dumerilii c-opsin going along with a corresponding gene structure. For r-opsins and tetraopsins gene structures are shown for A: Homo sapiens MELAN Q9UHM6, B: Apis mellifera UV opsin AAC47455.1, C: Limulus polyphemus opsin-5-like XP 013785122.1, and D: Homo sapiens OPN5 Q6U736. See Figure 2—figure supplement 1 for un-collapsed ML tree, Figure 2—figure supplement 2 for un-collapsed Phylobayes tree, Figure 2—figure supplement 3 for the whole set of gene structures, Figure 2—figure supplement 4 for intron phases, Figure 2—figure supplement 5 for an unrooted tree of only xenopsins, Figure 2—figure supplement 6 for a tree of only xenopsins plus a few c-opsins as outgroup, Figure 2—figure supplement 7 for a tree of xenopsins only plus a few cnidops as outgroup, Figure 2—figure supplement 8 for a tree of xenopsins only plus a few c-opsins and cnidops as outgroup and Figure 2—source data 1 for gene accession numbers.

Figure 2—source data 1

Accession numbers of the genes used for gene tree inference.

https://cdn.elifesciences.org/articles/55193/elife-55193-fig2-data1-v2.xlsx

Gene structure analysis corroborates molecular phylogeny

Several sequences (for example from Idiosepius paradoxus and Terebratalia transversa), which in Ramirez et al., 2016, Vöcking et al., 2017, Rawlinson et al., 2019 and this study group within xenopsins were earlier classified as c-opsins (Passamaneck et al., 2011; Yoshida et al., 2015). This view was either based on automated gene annotation, similarity searches, or phylogenetic analyses with only low taxon sampling. Nonetheless, it is in congruence with the presence of the NKQ tripeptide pattern (Figure 1) in the fourth cytoplasmic loop, which is in c-opsins crucial for specific binding to Gαi/t (Marin et al., 2000). To test if the grouping of the new opsin sequences found in T. inopinata and M. fuliginosus may result from tree inference artifacts — we cloned the respective genes from genomic DNA, analyzed gene structure and mapped it together with gene structure data generated by Vöcking et al., 2017 onto the protein alignment. Both the xenopsin and the c-opsin groups have specific gene structures. Three introns are highly conserved in position and intron phase throughout c-opsins. In comparison, two distinct introns in xenopsins are conserved likewise in position and intron phase (Figure 2, Figure 2—figure supplements 3 and 4). The gene structures of the new T. inopinata and M. fuliginosus xenopsins and the M. fuliginosus c-opsin match well with those of other xenopsins and c-opsins, respectively, and strongly corroborates the classification based on the molecular phylogenetic analysis. The xenopsin clade contains only sequences from protostomes. Notably, the closest related clade is neither a deuterostome specific nor a protostome specific opsin group, but cnidops. Yet, validation of this sister group relationship by gene structure data is not possible, since cnidops are lacking introns.

Xenopsin is expressed in cilia of the eye photoreceptor cells in larval T. inopinata

Larvae of T. inopinata possess one median eye apical of the anterior vibratile plume and one pair of lateral eyes halfway down from the apical to the abapical pole (Figure 3A). All eyes can be easily identified in live animals due to their red pigmentation. EM sections show that all three eyes form epidermal invaginations (Figure 4A, Figure 4—figure supplements 1A and 2), and whole-mount in situ hybridization revealed that Tin-xenopsin is strongly expressed in the region of all three eyes (Figure 3B,C). Besides, we found weak expression of Tin-xenopsin in few other cells, which are not associated with shielding pigments. One pair of cells lies on the rim of the anterior ciliary groove. Another pair lies lateral to the axial nerve running down from the apical organ (Figure 3B,C). These cells have small projections that also show weak expression of Tin-xenopsin (Figure 3—figure supplement 1A,B) and extend towards the body surface. A third pair lies lateral to the opening of the internal sac at the abapical pole (Figure 3B,C, Figure 3—figure supplement 1C). The in situ hybridization signal was much stronger in the eye regions than in the extraocular cells. Custom made antibodies against Tin-xenopsin specifically stain the eye regions (Figure 3D), but no significant staining appeared in the extraocular Tin-xenopsin expressing or any other cells.

Figure 3 with 1 supplement see all
Xenopsin expression in Tricellaria inopinata.

(A) Anterior view of a larva showing the pigment spots of the paired lateral eyes (filled arrowheads) and the single median eye (outlined arrowheads). (B,C) WMISH of Tin-xenopsin. Maximum projections of z-stacks spanning the whole larva. Single spots are labeled in the positions of the lateral eyes and two spots in the position of the single median eye. Several cells not associated with shielding pigment (asterisks) are also labeled. (D) Anti Tin-xenopsin antibody labels only the eyespot regions (filled yellow arrowheads: lateral eyes, outlined yellow arrowhead: median eye). (E,F) Combination of ISH and IHC. (E) Lateral eye. Tin-xenopsin antibody localizes adjacent to the mRNA around the nucleus of the eye photoreceptor cell (filled white arrowhead). (F) Median eye. Tin-xenopsin antibody localizes between a left and a right photoreceptor cell (outlined white arrowheads). See Figure 3—figure supplement 1 for details on Tin-xenopsin expression in extraocular cells.

Figure 4 with 3 supplements see all
Subcellular localization of xenopsin in the lateral eye of Tricellaria inopinata.

(A,B) Electron microscopic images (cryofixation) showing the photoreceptor cell (PRC) sending numerous cilia (ciPRC) into the eye invagination. The cilia possess basal bodies (white asterisks) and vertical (vr) and horizontal (hr) rootlets. Shielding pigment vesicles (black arrowheads) are present in the PRC and the adjacent pigmented coronal cells (PCC1, PCC2). Inset in B: cross-section of eye PRC cilia (chemical fixation) showing the 9 × 2 +2 organization of the axoneme. (C–F) IHC labeling of Tin-xenopsin and acetylated alpha-tubulin. Same orientation as in (A,B). Tin-xenopsin protein localizes within the cilia projecting into the eye invagination of the eye PRC. The basal bodies (white asterisks) are visible inside the eye PRC. ciBW: cilia of the body wall, cu: cuticle, nuPRC: nucleus of the photoreceptor cell. See Figure 4—figure supplement 1 for Tin-xenopsin localization in the median eye, Figure 4—figure supplement 2 for the cellular composition of the lateral eye, and Figure 4—figure supplement 3 for differences in the appearance of shielding pigment granules between chemical and cryofixation.

To get insights into the fine structure of the eyes, we performed serial section electron microscopy. The invagination of the lateral eyes is 5 μm deep, and it is formed by two neighboring coronal epidermal cells (PCC1 and PCC2 in Figure 4 and Figure 4—figure supplement 2) and the eye photoreceptor cell (PRC in Figure 4 and Figure 4—figure supplement 2). The coronal cells differ from adjacent coronal cells by the lack of cilia and the presence of abundant shielding pigment vesicles in the region of the eye invagination. The vesicles show high electron density in the chemically fixed specimen, but moderate electron density in the cryo-fixed specimen (Figure 4B, Figure 4—figure supplement 3). The two coronal cells line the apical and the lateral walls of the invagination, while the eye PRC lines the bottom and the abapical wall (Figure 4—figure supplement 2). The sensory cell bears a very dense bundle of cilia (ciPRC in Figure 4A,B; Figure 4—figure supplement 2) extending into the eye invagination. In the right eye of the cryo-fixed specimen, we counted 170 cilia. The axonemal microtubules of the cilia are arranged in 9 × 2+two pattern (Figure 4B). The cilia of the PRC penetrate the cuticle (cu in Figure 4A,B) and fill most of the eye invagination. The individual cilia have a diameter of 200 nm and are around 11 μm in length, and their upper halves extend above the body surface. Accordingly, the total surface of the ciliary membranes is approximately 1170 μm2. No other cell sends cilia into the invagination. The perikaryon of the eye PRC lies anterior to the base of the invagination (Figure 4—figure supplement 2). A single axon extends from the basal part of the sensory cell, joins the equatorial nerve ring, and runs towards the anterior ciliary groove. Abapical to the eye sensory cells lies an additional sensory cell (aSC in Figure 4—figure supplement 2). It sends a slender dendrite running upwards on the abapical side of the eye PRC and forms an anteriorly projecting pillar-like elevation emerging from the abapical wall of the eye invagination (Figure 4—figure supplement 2). On top of the elevation, 15 cilia with a 9 × 2 +two axoneme emerge from the tip of the dendrite and penetrate the cuticle.

The invagination of the median eye is formed by two coronal cells with shielding pigment granules and two PRCs (PRC1, PRC2 in Figure 4—figure supplement 1A). Subcellular characteristics of the coronal cells and PRCs are similar to those of the lateral eyes, but the cellular arrangement is different. The coronal cells line the bottom as well as the apical and abapical walls of the invagination, while the two photoreceptor cells line the left and the right wall. The perikarya of the PRCs lie distant to each other on the left and the right from the invagination and the ciliary bundles of the PRCs project from both sides into the eye invagination.

Knowing the ultrastructure of the eyes makes it possible to localize Tin-xenopsin mRNA and protein on the subcellular level by combining fluorescence in situ hybridization (FISH) with immunohistochemistry. In the lateral eye, the Tin-xenopsin FISH signal surrounds a nucleus next to the base of the eye invagination (Figure 3E). It matches well the position of the eye PRC nucleus in the EM dataset. The anti-Tin-xenopsin antibody signal is directly adjacent to the FISH signal and co-localizes with the cilia labeled by anti-acetylated α-tubulin in the eye invagination (Figure 4C–F). The median eye shows a similar pattern. While the FISH signal stains one cell on each side of the invagination, the opsin antibody stains the cilia inside the eye invagination (Figure 3F; Figure 4—figure supplement 1B–E). Accordingly, Tin-xenopsin mRNA is located throughout the soma of the eye photoreceptor cells of T. inopinata, whereas the opsin protein resides in the ciliary bundles emerging from these cells.

Tin-xenopsin likely is most sensitive in blue light and triggers the phototactic response of the larva

Since we could not detect the expression of any other opsin than Tin-xenopsin in the eyes of T. inopinata, we were interested in behavioral responses, which depend on directional detection of light by the eyes for the first functional characterization of this new opsin. We assayed the phototactic displacement of freshly hatched larvae under different wavelengths. The animals showed the biggest displacement towards blue light (454 nm) but still showed displacement towards green (513 nm), cyan (506 nm), and purple/UV (407 nm) (Figure 5, Figure 5—figure supplement 1). We could not detect a reaction with wavelength beyond the green spectrum (593 nm, 612 nm, 630 nm).

Figure 5 with 1 supplement see all
Spectral response of Tricellaria inopinata larvae.

(A) One-dimensional displacement of larvae during stimulation with blue (454 nm) light. Each recording started with no stimulus for 30 s. Afterwards, the light stimulus was activated for 15 s, followed by another 45 s in darkness. To generate violin plots, all tracked positions during a time of guaranteed illumination were used (seconds 40 to 42, dashed box). (B) Violin plot of the spectral response of the larvae. The animals show the greatest displacement under blue light (454 nm). Within the green and violet spectrum, the animals still respond positively, but further in the ultraviolet and wavelength beyond yellow (593 nm) only weak reactions were detectable. Violin plots based on videos containing between 50 to 230 animals each: 375 nm n = 5; 407 nm n = 4; 455 nm n = 5; 506 nm n = 8; 515 nm n = 13; 593 nm n = 4; 612 nm n = 3; 630 nm n = 3; Dark n = 3. See Figure 5—figure supplement 1 for violin plots of each individual experiment, Figure 5—source data 1 for raw data of graph in A and Figure 5—source data 2 for raw data for the graph in B.

Figure 5—source data 1

Raw data of behavioral experiment on larval displacement during stimulation with blue light.

https://cdn.elifesciences.org/articles/55193/elife-55193-fig5-data1-v2.xlsx
Figure 5—source data 2

Raw data of behavioral experiments on the spectral response of the larva.

https://cdn.elifesciences.org/articles/55193/elife-55193-fig5-data2-v2.xlsx

Xenopsin is coexpressed with r-opsin in cerebral eye PRCs in larval M. fuliginosus

M.M. fuliginosus larvae develop three pairs of pigmented eyespots in the head - one in a midventral position, one mediodorsal, and the third one in a laterodorsal position (Figure 6A,B). The ventral eyespots develop first at around 14 hpf, and the two pairs of dorsal eyespots develop at around 42 hpf. Preliminary investigation of ultrastructural data at 72 hpf stage revealed a mainly rhabdomeric organization of the ventral and mediodorsal eyespots, whereas the third laterodorsal eyespot revealed a ciliary structure. The ventral eyespot has three photoreceptor cells (PRCs), sending dense microvilli into the concavity made by two pigment cup cells (PCs) (Figure 6G–K, Figure 6—figure supplement 1). The PRCs are arranged adjacent to each other with the first PRC (PRC1) positioned medially, the second PRC (PRC2) in the middle, and the third PRC (PRC3) laterally. The dorsal rhabdomeric and ciliary eyespots are composed of one PRC and one PC.

Figure 6 with 1 supplement see all
Xenopsin in the dorsal and ventral eyes of Malacoceros fuliginosus.

(A,B) Light micrographs of ventral (arrowhead) and dorsal (asterisk) microvillar eyes at 48 hpf. (C,D) WMISH of Mfu-xenopsin in the ventral (arrowhead) and dorsal (black asterisk) eyes. (E–F’’’) Double FISH of Mfu-xenopsin and Mfu-r-opsin3. Mfu-xenopsin co-localizes with Mfu-r-opsin3 in all three PRCs of the ventral eye (E’’’). Numbers indicate the PRCs in the order of their development. (F–F’’’) Mfu-xenopsin and Mfu-r-opsin3 colocalization in the dorsal eye PRC (F’’’). Mfu-xenopsin is also expressed in an adjacent cell (white asterisk). (G–K) Ultrastructure of the second ventral eye photoreceptor cell (PRC2) depicting the cilium (ciPRC2, highlighted in green). (L–N) Antibody labeling against Mfu-r-opsin3 and acetylated alpha-tubulin reveals a prominent cilium emerging in between the r-opsin3+ microvilli in both the ventral and the dorsal eye. bb: basal body; mvPRC1: microvilli of PRC1; mvPRC2: microvilli of PRC2; PRC1: first PRC; smPRC1: submicrovillar cisternae of PRC1; smPRC2: submicrovillar cisternae of PRC2.

RNA in situ hybridization revealed the specific expression of Mfu-xenopsin in both dorsal and ventral rhabdomeric eyes but not in the ciliary eyes. Further, double FISH with Mfu-r-opsin3 (expressed in all rhabdomeric PRCs) confirmed Mfu-xenopsin expression in all three PRCs of the ventral eye (Figure 6E–E’’’). In dorsal rhabdomeric eyespot, however, in addition to its expression in the Mfu-r-opsin3+ PRC, we detected Mfu-xenopsin in an adjacent cell (Figure 6F–F’’’).

To assess the presence of ciliary structures in the ventral and dorsal eye and to achieve quantitative data on the surface extension of microvillar and ciliary structures, we analyzed a 3D electron microscopic data set of ventral and dorsal eyes in a 72 hpf stage larva in detail. In the lateral and the medial cells of the ventral eyes, only a basal body with an accessory centriole underneath the apical cell membrane (in the medial cell we could see an accessory centriole only on the right body side), but no cilia are present, which gives rise to the microvillar brushes (Figure 6—figure supplement 1). In contrast, the middle cell bears a long cilium projecting together with the microvillar brushes into the eye cavity (Figure 6G–K). This cilium is also visible in light microscopic stainings (Figure 6L). We estimated the microvillar surface of the middle PRC as 296 μm2 based on the average diameter of the microvilli, the number of microvilli per area, and the total volume of the space filled by the microvilli assessed from the 3D image stack. The ciliary surface is 10.7 μm2 based on the length and the diameter of the cilium. Accordingly, the ratio of ciliary to microvillar membrane surface is 1:27.7. The PRC of the dorsal eye likewise possesses a long cilium projecting together with the PRC microvilli into the eye cavity (Figure 6M–N).

Discussion

M. fuliginosus is the first organism known to have both xenopsin and c-opsin

Based on phylogenies with broad taxon sampling across the animal kingdom, Ramirez et al., 2016 and Vöcking et al., 2017 reported a secondary loss of xenopsins, as well as c-opsins in several major animal groups. Notably, xenopsins and c-opsins were not known to occur together. Annelids are the only group in which both opsin types were found, while other spiralians have only xenopsin and arthropods and deuterostomes have only c-opsins (Ramirez et al., 2016; Rawlinson et al., 2019; Vöcking et al., 2017). But even within annelids, mutually exclusive distribution of these opsins was reported. Xenopsin was only found in the basally branching oweniids (Vöcking et al., 2017), whereas c-opsins were only found in Platynereis dumerilii (Arendt et al., 2004) and sabellids (Bok et al., 2017) and genomic loss of both opsins are evident for Capitella teleta and Helobdella robusta (Vöcking et al., 2017). To our knowledge, not a single organism has been hitherto reported to have both a c-opsin and a xenopsin. No evolutionary or functional explanation has been given so far as to why xenopsins and c-opsins do not co-occur in any of the animals screened. Accordingly, some uncertainty remained, whether the distinction between xenopsins and c-opsins is a mere tree inference artefact. We now provide evidence for an independent origin of these opsins based on thorough molecular phylogeny, the exon-intron structure of the genes and a clear case of co-occurrence in a single species, M. fuliginosus.

Xenopsin is employed in the eyes of several protostomes

In vertebrates, c-opsins constitute the visual pigments of the retinal rods and cones in the eyes and serve additional functions when expressed in the pineal or deep brain PRCs (Blackshaw and Snyder, 1999; Kawano-Yamashita et al., 2014; Hankins et al., 2014). In protostomes, c-opsins were only reported from annelids and arthropods (Bok et al., 2017; Cronin and Porter, 2014; Hering and Mayer, 2014; Ramirez et al., 2016; Vöcking et al., 2017). Moreover, the expression of c-opsin has not been reported from PRCs in cerebral eyes, but from extraocular brain photoreceptors and in the case of the annelid subgroup of Sabellida in tentacular crown eyes (Arendt et al., 2004; Beckmann et al., 2015; Bok et al., 2017; Velarde et al., 2005; Verasztó et al., 2018). Instead, in many protostomes, r-opsins sense light in microvillar eye photoreceptor cells (Fain et al., 2010; Ramirez et al., 2016; Terakita, 2005). Besides, the evidence is increasing that xenopsins play important roles in protostome eyes. So far, xenopsin expression has been reported in the eye PRCs and serially homologous extraocular photoreceptor cells in chiton larva (Vöcking et al., 2017), in the eyes of larval brachiopods (Passamaneck et al., 2011) and recently in flatworm brain PRCs (Rawlinson et al., 2019). In this study, we report expression in the eyes of a larval annelid and eyes of a larval bryozoan. Seemingly, xenopsin is more common in the eyes of those protostomesthan hithero anticipated.

Xenopsin enters cilia

Subcellular targeting of opsins is an important prerequisite for visual perception in PRCs. Being transmembrane proteins, opsins travel integrated into vesicle membranes from the Golgi to the plasma membrane. Once there, they can enter plane plasma membrane areas similar to melanopsin in intrinsic light-sensitive retinal ganglion cells in the vertebrate retina (Belenky et al., 2003) and like many of the vertebrate non-visual c-opsins expressed in deep brain receptors, inner layers of the retina and in several other tissues (Foster and Bellingham, 2004; Hunt et al., 2014). Access to specialized membrane extensions like cilia or microvilli depends on specific active transport mechanisms (Schopf and Huber, 2017; Wang and Deretic, 2014; Wingfield et al., 2018). Though the evolution of sequence motifs relevant for protein binding to the respective transport machinery is not well understood, the capability of certain opsin types to enter either cilia or microvilli is seemingly very well conserved. To our knowledge, no c-opsin entering microvilli and no r-opsin entering cilia are known. We provide unambiguous evidence that xenopsin enters cilia in T. inopinata. Likely, this is also the case in the eye photoreceptor cells of the larval brachiopod Terebratalia, where xenopsin expressing cells show a ciliary organization (Passamaneck et al., 2011) and in brain PRCs in the flatworm Maritigrella (Rawlinson et al., 2019). The subcellular localization of xenopsin in PRCs of larval Leptochiton asellus (Vöcking et al., 2017) and larval M. fuliginosus is not clear since custom-made antibodies against the opsin did not reveal positive staining. However, in Leptochiton asellus, the presence of prominent ciliary structures beside microvilli expressing r-opsin provides the structural prerequisites for a similar opsin targeting. Similarly, in M. fuliginosus the dorsal eye PRC and the second ventral eye PRC bear a prominent cilium in addition to microvilli. Whether the xenopsin expressed enters these cilia remains speculative. Only for xenopsin A exist data on the subcellular localization of the protein (always in cilia), but so far not for xenopsin B. The first and third PRC of the ventral eye bear in addition to microvilli only basal bodies and accessory centriols close to the apical surface, which may be remnants of cilia. The xenopsin expressed in these cells may not enter any surface extensions, enter microvilli or may even not be translated into protein. If it would enter microvilli in certain PRCs of M. fuliginosus, xenopsin would be the first opsin group known to have the capability to enter both cilia and microvilli.

Evolution of bilaterian eye PRCs

For a long time, hypotheses on the evolution of eye PRCs focused mainly on PRCs expressing either r-opsin or c-opsin. R-opsin expressing cells employing the same kind of phototransduction cascade and with similar electrophysiological responses are found in the eyes of protostomes and deuterostomes (Arendt et al., 2002; Gomez et al., 2009; Fain et al., 2010; Koyanagi et al., 2005; Panda et al., 2002; Shichida and Matsuyama, 2009). Accordingly and due to conserved patterns in development, the presence of these PRCs already in the eyes of the bilaterian ancestor has been suggested (Arendt, 2003; Arendt, 2008; Arendt et al., 2004; Fernald, 2006; Gehring, 2014; Lamb, 2013; Shubin et al., 2009). Though c-opsins detect light in rods and cones of the vertebrate retina, its ancestral expression is assumed in brain extraocular photoreceptors (Arendt, 2008; Arendt et al., 2004; Shubin et al., 2009). Only in such cells were c-opsins found in arthropods (Beckmann et al., 2015; Velarde et al., 2005) and annelids (Arendt et al., 2004). Similarly, many kinds of non-visual c-opsins of vertebrates like encephalopsins, TMT-opsins, or VA-opsins are found in the brain in addition to possible functions in the inner layers of the retina (Hunt et al., 2014; Pérez et al., 2019). Accordingly, the employment of c-opsin cells in the visual cells of cerebral eyes evolved later, most probably in the lineage leading to chordates (Vopalensky et al., 2012).

Morphological data on the presence of ciliary PRCs in the eyes of several less studied protostome animals like bryozoans (Reed et al., 1988; Woollacott and Eakin, 1973; Woollacott and Zimmer, 1972), gastrotrichs (Woollacott and Eakin, 1973) and nemerteans (von Döhren and Bartolomaeus, 2018) are, however, in conflict with this scenario. Vöcking et al., 2017 proposed that in addition to rhabdomeric and c-opsin, xenopsin is a third important player in PRC and eye evolution questioning a common origin of ciliary eye PRCs in protostomes and deuterostomes. Our study is providing further support for this view.

Though in protostomes, c-opsins only exist in annelids and arthropods, ancestral employment in extraocular brain PRCs is still likely. But seemingly many kinds of protostome ciliary PRCs do not employ c-opsin. Instead, cellular expression of xenopsin is reported from ciliary PRCs in the eyes of larval brachiopods (Passamaneck et al., 2011) and bryozoans (this study) and meanwhile also from extraocular and eye ciliary PRCs in flatworms (Rawlinson et al., 2019). Further, the presence of xenopsin is also known from molluscs, annelids, chaetognaths, and rotifers (Ramirez et al., 2016; Rawlinson et al., 2019; Vöcking et al., 2017). Coexpression with r-opsin is evident in a larval chiton (Vöcking et al., 2017) and an annelid (this study) in microvillar eye PRCs, which partly also exhibit ciliary structures. Xenopsin may have been co-opted by these mainly microvillar cells (Figure 7 scenario A), but it is also conceivable that the observed cellular coexpression with r-opsin in two subgroups of lophotrochozoans points towards an evolutionary link between ciliary and microvillar PRCs. During the evolution of protostomes eyes, formerly microvillar PRCs may have changed into mixed microvillar/ciliary cells coexpressing r-opsin and xenopsin (Figure 7, scenario B). Since co-expression is evident from xenopsin A and B, this happened likely before the diversification of xenopsins in protostomes. Clear hypotheses on the ancestral targeting of xenopsin (cilia and/or microvilli) may need further investigation, but existing data so far point towards cilia as targets. Even the presence of mixed cells in the eyes of the last common ancestor of bilaterians is conceivable (Figure 7, scenario C), since opsin tree topology suggests a genomic loss of xenopsin in the deuterostome stem lineage. Within protostomes, the mixed organization was retained in some extant organisms (several molluscs, certain annelids) and transformed in other organisms into a purely ciliary or microvillar organization going along with loss or downregulation of r-opsins (bryozoans, brachiopods) or xenopsins (arthropods, certain annelids), respectively. Such a hypothesis also raises the question, whether the co-occurrence of ciliary and microvillar PRCs within the same eye as known, for example from several molluscs (Bartolomaeus, 1992; Blumer, 1998; Salvini-Plawen, 2008) or the larva of the polyclad flatworms (Eakin and Brandenburger, 1981; Rawlinson et al., 2019) may be the result of integrating or co-opting ciliary cells into microvillar eyes or caused by duplication and diversification of formerly mixed microvillar/ciliary PRCs. Interestingly, the polyclad flatworm Maritigrella crozieri has several kinds of ciliary PRCs, which are expressing xenopsin in adults outside of pigmented eyes, in larval epidermal eyes, and in larval cerebral eyes, which also contain r-opsin expressing microvillar PRCs (Rawlinson et al., 2019). The evolutionary origin of the extraocular PRCs and the epidermal eyes is unclear. Nonetheless, developmental data from Schmidtea mediterranea, indicate homology of flatworm cerebral eyes to those of other protostomes (Lapan and Reddien, 2011; Lapan and Reddien, 2012). Therefore, origination from eyes with mixed r-opsin/xenopsin+ microvillar/ciliary PRCs is conceivable.

Scenarios on eye PRC evolution in Bilateria.

The bilaterian ancestor had extraocular c-opsin+ ciliary PRCs. These became integrated into the eyes in the lineage leading to vertebrates and were lost in many protostomes along with secondary loss of c-opsin. Scenario A: Cerebral eyes contained microvillar r-opsin+ PRCs in the bilaterian ancestor. Xenopsin was co-opted several times independently by microvillar PRCs, and eye PRCs were several times independently transformed into or replaced by ciliary xenopsin+ PRCs. Scenario B: Ancestral r-opsin+ microvillar eye PRCs were transformed once into mixed microvillar/ciliary PRCs coexpressing r-opsin and xenopsin. In extant organisms, those were retained or changed into purely microvillar r-opsin+ or ciliary xenopsin+ PRCs going along with genomic loss or downregulation of xenopsin or r-opsin, respectively. Scenario C: Mixed ciliary/microvillar PRCs were already present in the bilaterian ancestor.

Xenopsin function and physiology

Strong phototactic responses, as we observed in the larva of T. inopinata, depend on directional detection of light by pigmented eyes. Since we could not find any opsin other than xenopsin expressed in the eyes of T. inopinata, we suggest that the xenopsin here is responsible for light reception and triggers the phototactic behavior of the larva. Accordingly, it has a similar visual function as r-opsins have in microvillar eye PRCs of several other protostomes (Fain et al., 2010; Jékely et al., 2008; Neal et al., 2019; Randel et al., 2013). The ciliary surface of the PRC in T. inopinata is even three times larger than the microvillar surface of the middle eye PRC in M. fuliginosus. Notably, heterologous expression of Maritigrella crozieri xenopsin suggests that it acts mainly via Gαi and possibly to a lower extent also via Gαs, but not via Gαq signaling (Rawlinson et al., 2019) as r-opsins do (Fain et al., 2010; Shichida and Matsuyama, 2009). Hence, similar to the case of r-opsins and c-opsins, downstream signaling and even the cellular electrophysiological response may also be different upon the excitation of r-opsins and xenopsins.

Most PRCs express only one type of visual opsin. If PRCs employ more than one opsin, they are usually from the same subgroup of visual pigments and activate the same transduction cascade (Applebury et al., 2000; Arikawa et al., 2003; Dalton et al., 2015; Isayama et al., 2014; Katti et al., 2010; Parry and Bowmaker, 2016; Rajkumar et al., 2010), which is suggested, ultimately, to expand the visual spectrum. The same function is assumed in the annelid P. dumerilii, where Go-opsin and r-opsin co-occur in eye PRCs (Gühmann et al., 2015). If Gαi signaling of xenopsin is conserved in protostomes, PRCs coexpressing xenopsin and r-opsin, as we observed in M. fuliginosus and are known in Leptochiton asellus (Vöcking et al., 2017), might potentially be polymodal sensory cells with complex physiology capable of integrating different stimuli by activation of different signaling cascades. The ciliary surface of the middle eye PRC of M. fuliginosus is nearly 30 times less than the microvillar surface. This may hint to a minor role of xenopsin signaling in this eye, but other parameters like the efficiency of the specific sensory transduction pathway and the protein content in the membrane certainly impact the sensitivity as well. The contribution of cilia in light detection may be higher in eyes where microvilli are accompanied by higher numbers of cilia as described in several molluscs (Blumer, 1995; Blumer, 1996; Hughes, 1970; Zhukov et al., 2006).

Conclusion

Xenopsin seems to be an important visual pigment in protostome eyes. This opsin type was overlooked for a long time, probably because most molecular data on protostome light perception are from arthropods, which secondarily lost xenopsin. M. fuliginosus is the first organism known to have xenopsin and c-opsin corroborating the distinct evolutionary origin of these opsin types inferred from phylogenetic and gene structure analysis. All other organisms studied thus far, for reasons unknown, have either c-opsin or xenopsin. Xenopsin, like c-opsin, enters cilia. In protostomes, it is employed in purely ciliary PRCs and found coexpressed with r-opsin in microvillar PRCs that also have ciliary structures. Xenopsin or xenopsin expressing cells might have been recruited several times independently in the eyes of protostomes. Alternatively, these eyes already early in evolution employed a possibly polymodal r-opsin+ and xenopsin+ microvillar/ciliary PRCs. Further studies on the employment of xenopsin in protostomes will be of high interest for a better understanding of evolution, function, and plasticity of animal photoreceptor cells and eyes. Counterparts of vertebrate c-opsin employing ciliary PRCs in protostomes probably exist only in certain annelids and arthropods, since c-opsin according to available sequence resources has been lost in all other protostome animals.

Materials and methods

Key resources table
Reagent type
(species) or resource
DesignationSource or referenceIdentifiersAdditional information
Gene (Tricellaria inopinata)XenopsinGenbankMT901641
Gene (Malacoceros fuliginosus)XenopsinGenbankMT901639
Gene (Malacoceros fuliginosus)Ciliary opsinGenbankMT901640
Strain, strain background (Malacoceros fuliginosus)Wild typeUniversity of Bergen, Sars Centre for Marine Molecular BiologyNCBITaxon: 271776
Antibodymouse monoclonal anti-acetylated α-tubulin IgG1Sigma-AldrichRRID:AB_609894Dilution
1:300 (Mfu)
1:50 (Tin)
AntibodyRat polyclonal anti-Mfu-r-opsin3 IgGUniversity of Bergen, Sars Centre for Marine Molecular Biology1:100
AntibodyRabit polyclonal anti-Tin-xenopsin IgGUniversity of Bergen, Sars Centre for Marine Molecular Biology1:500
AntibodyAlexa Fluor 633 goat monoclonal anti-rat IgGThermoFisher ScientificRRID:AB_25357491:500
AntibodyAlexa Fluor 488 goat momoclonal anti-mouse IgGThermoFisher ScientificRRID:AB_25357641:500
Recombinant
DNA reagent
PGem-T-Tin-xenops (plasmid)University of Bergen, Sars Centre for Marine Molecular BiologyUsed for synthesizing WMISH probes
Recombinant
DNA reagent
PGem-T-Mfu-xenops (plasmid)University of Bergen, Sars Centre for Marine Molecular BiologyUsed for synthesizing WMISH probes
Recombinant
DNA reagent
PGem-T-Mfu-cops (plasmid)University of Bergen, Sars Centre for Marine Molecular BiologyUsed for synthesizing WMISH probes
Sequence-based reagentMfu-xenops-WMISH forward primer (5’- > 3’)University of Bergen, Sars Centre for Marine Molecular Biology5’-CACCATCATGTTGAATAATGACTCCTACTC-3’
Sequence-based reagentMfu-xenops-WMISH reverse primer (5’- > 3’)University of Bergen, Sars Centre for Marine Molecular Biology5’-GATTCGTGGAATGCTGATTTGTGAC-3’
Sequence-based reagentMfu-cops-WMISH forward primer (5’- > 3’)University of Bergen, Sars Centre for Marine Molecular Biology5’-ATCACACAGGATATCACAAATGCCTCAG-3’
Sequence-based reagentMfu-cops-WMISH reverse primer (5’- > 3’)University of Bergen, Sars Centre for Marine Molecular Biology5’-GCAATAACGATGTCACCTGGACATTG-3’
Sequence-based reagentTin_xenopsin-WMISH forward primer (5’- > 3’)University of Bergen, Sars Centre for Marine Molecular Biology5’-CTTATGGTCATTGCTGT-3’
Sequence-based reagentTin_xenopsin-WMISH reverse primer (5’- > 3’)University of Bergen, Sars Centre for Marine Molecular Biology5’-CACCCTGCCATTAGTC-3’
Sequence-based reagentTin_xenopsin-WMISH forward nested primer (5’- > 3’)University of Bergen, Sars Centre for Marine Molecular Biology5’-TGGGGGTTGTTTTGGTCGT-3’
Sequence-based reagentTin_xenopsin-WMISH reverse nested primer (5’- > 3’)University of Bergen, Sars Centre for Marine Molecular Biology5’-CTGTTGCCTTCTTCTCTCGT-3’
Commercial assay or kitSuperscript III First-Strand Synthesis SystemThermoFisher ScientificCatalog number: 18080051
Commercial assay or kitRNeasy Mini Kit
QiagenCatalog number: 74104
Software, algorithmIQ-TREEhttp://www.iqtree.org/RRID:SCR_017254
Software, algorithmPhylobayes-MPIhttps://github.com/bayesiancook/pbmpiRRID:SCR_006402
Software, algorithmMAFFT 7https://mafft.cbrc.jp/alignment/server/RRID:SCR_011811
Software, algorithmCLC Main WorkstationQiagenRRID:SCR_000354
Software, algorithmImageJNIHRRID:SCR_003070
Software, algorithmImaris 8.41BitplaneRRID:SCR_007370

Animal culture

Request a detailed protocol

Adults of the polychaete Malacoceros fuliginosus (Claparède, 1868) were collected from Pointe de Mousterlin, Fouesnant, France. The animals were maintained in the lab facility in sediment containing seawater tanks at 18°C and fed with ground fish food flakes (TetraMin granules, Tetra). Mature males and females were picked, rinsed several times with filtered seawater, and kept in separate bowls until they spawned. Staging was started from the time gametes were combined in a fresh bowl. Bowls were kept at 18°C under 12:12 hr light-dark cycle, and water was replaced every day or every second day. Larvae were fed with the microalga Chaetoceros calcitrans from 24 hpf onwards after each water change. Colonies of the bryozoan Tricellaria inopinata (d'Hondt & Occhipinti Ambrogi, 1985) were collected in Brest, France (48°23’38.3“N 4°25’57.4“W). The colonies were maintained at 18°C under 12:12 hr light-dark cycle in the lab animal facility.

RNA-seq and transcriptome assembly

Request a detailed protocol

For studies on M. fuliginosus, we used transcriptomic resources prepared in an earlier study (Kumar et al., 2020) from pooled larvae of several stages. For T. inopinata, we performed RNA-seq and de novo transcriptome assembly. The release of larvae was triggered by the onset of light in the tanks. Two hours later, swimming larvae were attracted by a light bulb and cryo-fixed. We extracted total RNA using the Agencourt RNAdvance Tissue Kit (Beckman Coulter, Brea, California). Library preparation and sequencing were performed by EMBL (Heidelberg, Germany) Genomics Core Facility using cation-based chemical fragmentation of RNA, Illumina (San Diego, California) Truseq RNA-Sample Preparation Kit and 1 lane of 100 bp paired-end read sequencing on Illumina HiSeq 2000. We used Cutadapt 1.2.1 (RRID:SCR_011841) for trimming and the ErrorCorrectReads tool implemented in Allpaths-LG (RRID:SCR_010742) for error correction of the raw reads and Trinity (RRID:SCR_013048) for de novo assembly. We performed two rounds of RNA-seq and assembly. For the first data set (assembly 1) the collected colonies were thoroughly cleaned. However, by microscopic inspection, we observed a minor proportion of zooids of other bryozoan species, which were dispersed across the colonies and could not be entirely removed. We assessed contamination of the respective assembly by screening for cytochrome oxidase subunit I (COI), and we found sequences indeed from four different bryozoans, but none from other animal groups. The second data set (assembly 2) showed contamination also with sequences from other taxa and was only used for corroboration and elongation of sequences retrieved from assembly 1.

Opsin tree inference

Request a detailed protocol

Retrieved opsin sequences from T. inopinata and M. fuliginosus were added to the set of opsin sequences (https://doi.org/10.7554/eLife.23435.009) analyzed by Vöcking et al., 2017. Sequences were first aligned with MAFFT 7 (RRID:SCR_011811) with option E-INS-I, ambiguously aligned N- and C-terminal regions were trimmed, sequences shorter than 160 amino acids removed and the remaining set of sequences again aligned with MAFFT seven with option E-INS-I. The output was manually edited to remove gap rich regions and ambiguously aligned positions. Maximum-likelihood phylogenetic analyses were run with IQ-TREE 1.5.5 (RRID:SCR_017254) with model LG+F+R8 chosen by ModelFinder, SH-like approximate likelihood ratio test (1000 replicates), ultrafast bootstrap (1000 replicates) and approximate Bayes test for estimating branch support, unsuccessful iterations to stop tree searching set to 500 and perturbation strength to 0.2. Bayesian analysis was performed with Phylobayes-MPI 1.6 (RRID:SCR_006402) running three chains for 90.000 cycles using the dataset specific substitution matrix (DS-GTR) generated by Vöcking et al., 2017 with parametric Γ modeling. Phylogenetic convergence of the chains was assessed with bpcomp.

Opsin gene structure analysis

Request a detailed protocol

For the two opsins found in M. fuliginosus and the one found in T. inopinata, the whole genes were cloned from genomic DNA for subsequent analysis of exon-intron boundaries. Genomic DNA was extracted with the Nucleospin Tissue Kit (Machery-Nagel, Düren, Germany) and tested for fragment length larger than 20 kb. As a starting point, gene-specific primers were designed based on the transcript sequences. For genome walking, four libraries were prepared with Universal Genome Walker Kit (Takara Bio, Mountain View, California) by enzymatic digestion and used for sequence elongation starting from exonic fragments. In parallel, long amplicons bridging smaller introns were also directly amplified from genomic DNA using LA Taq (Takara Bio), iProof (Biorad, Hercules, California, USA), and HotStarTaq Plus (Qiagen, Hilden, Germany) polymerases. Obtained amplicons up to 8 kb were cloned using pGEM-T easy Vector (Promega, Madison, Wisconsin) TOPO XL PCR cloning kit (Thermo Fisher Scientific), TopTen chemically competent cells (Thermo Fisher Scientific, Waltham, Massachusetts) and Sanger sequenced. Obtained sequences were used to design further primers for ongoing sequence elongation. Read assembly was performed with CLC Main Workstation (RRID:SCR_000354) 7.1. Gene structures of the opsins from M. fuliginosus and T. inopinata were determined based on the cloned genomic and the protein sequences retrieved from RNA-seq using WebScipio (Hatje et al., 2011). The obtained gene structures were together with the gene structures prepared by Vöcking et al., 2017 mapped onto the un-curated sequence alignment using Genepainter (Hammesfahr et al., 2013) to identify conservation of intron positions and phases.

Custom antibodies

Request a detailed protocol

Custom polyclonal antibodies were prepared and affinity-purified against peptides of Tin-xenopsin by 21st Century Biochemicals (Marlboro, Massachusetts) and Mfu-r-opsin-3 by Eurogentec (Liège, Belgium). For Tin-xenopsin, the peptide sequences VKAAGKKFGGDDAASQ from the 3rd cytoplasmic loop and ATKPAPSATQAPREKKATAL from the cytosolic tail and for Mfu-ropsin3 RHSEVPSGDGKKDTL and CKNRAIDKGGDEESDN both from the cytoplasmic tail were chosen as antigens. To assure antigen specificity, all peptide sequences were blasted against the T. inopinata, and M. fuliginosus transcriptome and gave only the respective opsin sequences as hits. The antibodies raised against the peptides ATKPAPSATQAPREKKATAL and RHSEVPSGDGKKDTL gave the best results and were used for the stainings.

Immunohistochemistry

Request a detailed protocol

M. fuliginosus larvae were first relaxed with 1:1 MgCl2-seawater for 3–5 min before fixing them in 4% PFA (in 1X PBS, 0.1% Tween20) for 30 min at RT. After fixing, the samples were washed two times in PTW followed by two washes in THT (0.1 M Tris pH 8.5, 0.1% Tween20). Blocking was in 5% sheep serum in THT for 1 hr before incubating in primary antibodies rat anti-Mfu r-opsin3, 1:100; monoclonal mouse anti-acetylated α-tubulin IgG (RRID:AB_609894) from Sigma Aldrich (Saint Louis, Missouri) 1:300 (Mfu), 1:50 (Tin), anti-Tin-xenopsin, 1: 500) for 48 hr at 4°C. The samples were then subjected to two 10 min washes in 1 M NaCl in THT followed by five 30 min washes in THT before incubating in secondary antibodies (Alexa Fluor 633 goat anti-rat IgG (RRID:AB_2535749), 1:500 and Alexa Fluor 488 goat anti-mouse IgG (RRID:AB_2535764), 1:500 (Thermo Fisher Scientific) overnight at 4°C. Next, the samples were washed in THT, two 5 min washes followed by five 30 min washes. Specimens were stored in embedding medium (90% glycerol, 1x PBS and 0.25% DABCO) at 4°C.

In situ hybridization

Request a detailed protocol

For gene cloning cDNA, was prepared from total RNA with Super Script II (Thermo Fisher Scientific), sequences of interest were PCR amplified with gene-specific primers, and amplicons were subsequently ligated into pgemT-easy vector (Thermo Fisher Scientific) and cloned into Top10 chemically competent E. coli (Thermo Fisher Scientific). Sanger sequencing was used to verify the cloned sequences before DIG- and FITC-labeled sense and antisense probes were generated with T7 and SP6-RNA Polymerases (Roche, Basel, Switzerland) or with Megascript Kit (Thermo Fisher Scientific). If needed, Smarter Race (Takara Bio) was used to elongate ends of transcript sequences. In situ hybridization experiments were performed as described previously (Vöcking et al., 2015) if formamide based hybridization buffers were used. Otherwise, we followed # (Sinigaglia et al., 2018) # for urea-based hybridization buffers. In brief, animals were fixed in 4% PFA in phosphate buffer and with Tween20 (PTW; pH 7.4) and subsequently washed and stored in methanol. For Tricellaria inopinata larvae, a 2 min prefixation with 0.3% Glutaraldehyde in 4% PFA was necessary. After rehydration in PTW, samples were briefly digested with Proteinase K, washed and prehybridized in hybridization with or without 5% dextran. Samples were hybridized with RNA probes for 72 hr. Tricellaria inopinata larvae required that each washing step after hybridization was extended to 30 min. Stainings were done with a combination of FastBlue (Sigma-Aldrich) and Fast Red (Roche). The significance of expression signals was evaluated by using sense probes as control experiments. All in situ hybridization experiments were performed on at least 15 specimens per gene for each sense, and anti-sense probe and the experiments were repeated at least two times.

Light microscopy

Request a detailed protocol

Light microscopic images were taken using Eclipse E800 (Nikon, Tokyo, Japan) and Examiner A.1 (Zeiss, Oberkochen, Germany), and confocal images were taken with an SP5 confocal microscope (Leica, Wetzlar, Germany). Image stacks were processed with ImageJ (RRID:SCR_003070), Imaris (RRID:SCR_007370) and Adobe Photoshop CC (RRID:SCR_014199).

Electron microscopy

Request a detailed protocol

For electron microscopic studies, two kinds of sample preparations were used. For chemical fixation larvae were relaxed for 3 min in 7% MgCl2 and seawater mixed 1:1 and then fixed in 2.5% glutaraldehyde in PBS, postfixed in 1% Osmium tetroxide in the same buffer, en-bloc stained with reduced Osmium, dehydrated in a graded ethanol series and embedded in Epon/Araldite as described in Vöcking et al., 2015. Cryo-fixation was performed at the EM Core facility at EMBL (Heidelberg, Germany). Larvae were relaxed as described above and then high-pressure-frozen with hexadecene acting as filler in an HPM 010 from Balzers (Balzers, Liechtenstein). Freeze substitution with 2% OsO4 and 0.1% uranyl acetate in a mixture of 95% acetone and 5% water was performed in a Leica (Wetzlar, Germany) EM AFS2 for 46 hr at −90°C. Samples were slowly warmed to −30°C, kept at this temperature for 6 hr, and slowly warmed to 0°C before they were taken out from the freeze-substitution device. Samples were rinsed several times in acetone at 0°C and at room temperature, stepwise transferred to Epon, and cured for 48 hr at 60°C.

Serial sections of 70 nm were cut with an ultra 35° diamond knife (Diatome, Biel, Switzerland) on a UC7 ultramicrotome (Leica) and collected on Beryllium-Copper slot grids (Synaptek, Reston, Virginia, USA) coated with Pioloform (Ted Pella, Redding, California, USA) and counterstained with 2% uranyl acetate and lead citrate. Complete series were imaged with STEM-in-SEM as described by Kuwajima et al., 2013 at a resolution of 4 nm/ pixel with a Supra 55VP (Zeiss, Oberkochen, Germany) equipped with Atlas (Zeiss) for automated large field of view imaging. Acquired images were processed with Adobe Photoshop CC, first registered rigidly followed by affine and elastic alignment (Saalfeld et al., 2012) with TrakEM2 (Cardona et al., 2012) implemented in Fiji (RRID:SCR_002285).

Behavioral assays of T. inopinata larva

Request a detailed protocol

Freshly hatched larvae were placed in a small clear plastic container that was situated inside of a chamber with two infrared long-pass filters (RG610, Reichmann Feinoptic, Brokdorf, Germany) installed above and below the container. The chamber was placed on a Zeiss Stemi 2000 stereo microscope. Through a hole on one side of the chamber, an LED served as the light stimulus. Eight different LEDs covered wavelengths from UV 375 nm to 630 nm in the red part of the spectrum. For each experiment between 50 to 230 freshly hatched larvae were placed inside the chamber. We recorded the reaction of the animals to the stimulus with an industrial monochrome CMOS camera (DMK 23U445, The Imaging Source, Bremen, Germany). Each recording starts in darkness for 30 s, followed by 15 s of illumination and another 45 s of darkness. The response of the animals to each wavelength was assayed at least three times, always with a new batch of animals and an extra four without any light stimulus as a control. From each recording, we removed the background and enhanced the contrast in Fiji. Subsequently, we tracked the animals' position in enhanced recordings for each frame with the Fiji plugin Trackmate (Tinevez et al., 2017). The tracking information was used to calculate the mean and median position of the animals for each frame for a single axis. To make different recordings comparable, we used the mean and median position of animals during the initial darkness to subtract from each position for each frame. Boxplots Violinplots were inferred from all the tracked positions of all the animals during a time of guaranteed illumination (second 40 to 42, Figure 5A, dashed box).

Data availability

Sequencing data have been deposited in Genbank under accession codes MT901639, MT901640, and MT901641. Source data files have been provided for Figures 2 and 5.

The following data sets were generated
    1. Doering CC
    2. Kumar S
    3. Tumu S
    4. Kourtesis I
    5. Hausen H
    (2020) NCBI GenBank
    ID MT901639. Malacoceros fuliginosus xenopsin gene, partial cds.
    1. Doering CC
    2. Kumar S
    3. Tumu S
    4. Kourtesis I
    5. Hausen H
    (2020) NCBI GenBank
    ID MT901640. Malacoceros fuliginosus ciliary opsin gene, complete cds.
    1. Doering CC
    2. Kumar S
    3. Tumu S
    4. Kourtesis I
    5. Hausen H
    (2020) NCBI GenBank
    ID MT901641. Tricellaria inopinata xenopsin gene, partial cds.

References

    1. Arendt D
    2. Tessmar K
    3. de Campos-Baptista MI
    4. Dorresteijn A
    5. Wittbrodt J
    (2002)
    Development of pigment-cup eyes in the polychaete Platynereis dumerilii and evolutionary conservation of larval eyes in bilateria
    Development 129:1143–1154.
    1. Arendt D
    (2003) Evolution of eyes and photoreceptor cell types
    The International Journal of Developmental Biology 47:563–571.
    https://doi.org/10.1387/IJDB.14756332
    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.
    1. Claparède E
    (1868)
    Les annélides chétopodes Du Golfe de Naples. Ramboz et schuchardt
    Genève.
  1. 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. Springer. pp. 105–135.
    https://doi.org/10.1007/978-1-4614-4355-1_4
  2. Book
    1. Eakin RM
    (1963)
    Lines of Evolution of Photoreceptors
    In: Mazia D, Tyler A, editors. General Physiology of Cell Specialization. McGraw-Hill. pp. 393–425.
    1. Gehring WJ
    (2014) The evolution of vision
    Wiley Interdisciplinary Reviews: Developmental Biology 3:1–40.
    https://doi.org/10.1002/wdev.96
  3. Book
    1. Hankins MW
    2. Davies WIL
    3. Foster RG
    (2014) The Evolution of Non-visual Photopigments in the Central Nervous System of Vertebrates
    In: Hunt D. M, Hankins M. W, Collin S. P, Marshall N. J, editors. Evolution of Visual and Non-Visual Pigments. Springer. pp. 105–135.
    https://doi.org/10.1007/978-1-4614-4355-1_3
    1. Hughes HPI
    (1970) A light and electron microscope study of some opisthobranch eyes
    Zeitschrift FüR Zellforschung Und Mikroskopische Anatomie 106:79–98.
    https://doi.org/10.1007/BF01027719
  4. Book
    1. Kawano-Yamashita E
    2. Koyanagi M
    3. Terakita A
    (2014) The Evolution and Diversity of Pineal and Parapineal Photopigments
    In: Hunt D. M, Hankins M. W, Collin S. P, Marshall N. J, editors. Evolution of Visual and Non-Visual Pigments. Springer. pp. 1–21.
    https://doi.org/10.1007/978-1-4614-4355-1_1
    1. Parry JWL
    2. Bowmaker JK
    (2016)
    Visual pigment coexpression in guinea pig cones : a microspectrophotometric study
    Investigative Ophthalmology & Visual Science 43:1662–1665.
    1. Shichida Y
    2. Matsuyama T
    (2009) Evolution of opsins and phototransduction
    Philosophical Transactions of the Royal Society B: Biological Sciences 364:2881–2895.
    https://doi.org/10.1098/rstb.2009.0051

Decision letter

  1. Dan Larhammar
    Reviewing Editor; Uppsala University, Sweden
  2. Diethard Tautz
    Senior Editor; Max-Planck Institute for Evolutionary Biology, Germany

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

Light receptor proteins in eyes come in three major types of which ciliary opsin and rhabdomeric opsin coexist in some animal groups including vertebrates. The newly discovered xenopsin has been reported to be expressed alone or coexist with rhabdomeric opsin. This report describes the first organism with all three opsin categories, the marine annelid Malacoceros fuliginosus.

Decision letter after peer review:

Thank you for submitting your article "The visual pigment xenopsin is widespread in protostome eyes and impacts the view on eye evolution" for consideration by eLife. Your article has been reviewed by two peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Diethard Tautz as the Senior Editor. The reviewers have opted to remain anonymous.

The Reviewing Editor has drafted this decision to help you prepare a revised submission.

Summary:

This is an interesting manuscript as it seems to resolve an important matter in visual opsin evolution by demonstrating presence of the enigmatic xenopsin together with c-opsin in an annelid, leading to the conclusion that xenopsin has evolved in parallel with the other two visual opsins (c-opsins and r-opsins). The study also shows that xenopsin localizes to larval cilia. Thereby it adds to a broader understanding of opsin evolution, eye evolution and the basis of visually guided behavior.

Essential revisions:

Generally, the reviewers found the manuscript well written and data carefully collected and well supported. They specifically commented that ISH and antibody stainings are of high quality and with beautiful transmission electron micrographs.

As an exception, the description of the behavioural assay did not clearly say whether a single experiment (with 50 to 100 animals) was performed, and then each animal's position taken as a data point, or whether the figure builds on multiple (how many?) experiments for each wavelength. Please be aware that the position of 100 animals all released together cannot be treated as independent data points; many if not the majority of such organisms do one or more of the following: avoid each other, follow each other, chemically communicate with each other. Thus, if the figure builds on just one of these experiments per wavelength, the data may not be treated correctly. There is little doubt that there is a reaction, but it would be good (and likely improve the data set) if the experiment was repeated several times with each wavelength, and the mean etc. of these separate experiments was presented.

Figure 2: The evidence for xenopsins and cnidopsins forming a clade still finds low to moderate support only and might therefore represent an artifact. The possibility remains that Xenopsins exist in Lophotrochozoans only and this should be mentioned and visualized in the model.

Even though many nodes in the xenopsin tree are well supported, it seems to suggest the existence of two paralogs in Lophotrochozoa that are distinctly present in molluscs, annelids and platyhelminths. This duplication appears to have occurred at the Lophotrochozoan root. If so, then the ciliary location would only be shown for one representative of one of the paralogs. This is important and should be mentioned.

For Malacoceros xenopsin, the reasoning that xenopsin locates to cilia does not work out because the two rhabdomeric photoreceptors that strongly express it do not have cilia (only basal bodies). Where would the opsin go? There is no evidence for 'rudimentary cilia', they are simply absent (no acetylated tubulin staining). This should be stated as is.

The figures, supplementary figures and figure legends seem to have been mixed in the new version as compared to the previous version. The numbers had disappeared. This confusing mess must be sorted out.

https://doi.org/10.7554/eLife.55193.sa1

Author response

Essential revisions:

Generally, the reviewers found the manuscript well written and data carefully collected and well supported. They specifically commented that ISH and antibody stainings are of high quality and with beautiful transmission electron micrographs.

As an exception, the description of the behavioural assay did not clearly say whether a single experiment (with 50 to 100 animals) was performed, and then each animal's position taken as a data point, or whether the figure builds on multiple (how many?) experiments for each wavelength. Please be aware that the position of 100 animals all released together cannot be treated as independent data points; many if not the majority of such organisms do one or more of the following: avoid each other, follow each other, chemically communicate with each other. Thus, if the figure builds on just one of these experiments per wavelength, the data may not be treated correctly. There is little doubt that there is a reaction, but it would be good (and likely improve the data set) if the experiment was repeated several times with each wavelength, and the mean etc. of these separate experiments was presented.

We agree that interactions of the larvae may introduce some bias and thus the new Figure 5 is now based on data from several independent experiments. For each wavelength we performed 3 up to 13 recordings and for each recording a new batch of 50 to 230 larvae was used. For Figure 5 we pooled for each wavelength all data obtained and performed descriptive statistics (violin plots) on the displacement of the larva. In the new Figure 5—figure supplement 1 we provide the plots for all individual experiments. Both Figure 5 and Figure 5—figure supplement 1 provide evidence for a clear phototactic reaction.

Main changes in the text are in the subsection “Behavioral assays of T. inopinata larva”. Figure 5, Figure 5—figure supplement 1 and the respective figure legends are updated.

Figure 2: The evidence for xenopsins and cnidopsins forming a clade still finds low to moderate support only and might therefore represent an artifact. The possibility remains that Xenopsins exist in Lophotrochozoans only and this should be mentioned and visualized in the model.

Even though many nodes in the xenopsin tree are well supported, it seems to suggest the existence of two paralogs in Lophotrochozoa that are distinctly present in molluscs, annelids and platyhelminths. This duplication appears to have occurred at the Lophotrochozoan root. If so, then the ciliary location would only be shown for one representative of one of the paralogs. This is important and should be mentioned.

To address the comments related to opsin evolution – the relation of xenopsins and cnidops and the diversification of xenopsins – we rerun the opsin tree with an improved taxon sampling by adding the recently published xenopsin sequences from the molluscs, Sepia officinalis and Ambigolimax valentianus, the bryozoan Bugula neritina, the flatworm Maritigrella crozieri, and the chaetognath Pterosagitta draco.

Figure 2 and all figure supplements of Figure 2 were updated and are based on the new inferred trees. Figure 2—figure supplements 5-8 are new and show trees of only xenopsins with different small outgroups. Main changes in the text are in the subsections “Behavioral assays of T. inopinata larva”, “Xenopsin is expressed in cilia of the eye photoreceptor cells in larval T. inopinata”, and “Evolution of bilaterian eye PRCs”.

In the new maximum-likelihood tree (IQ-TREE), for the clade composed of protostome xenopsins and cnidops, we find support values of ultrafast bootstrap and SH-like approximate likelihood ratio test ≥ 0.9 and approximate Bayes test ≥ 0.98. In a parallel Bayesian (Phylobayes) analysis this clade is supported with a posterior probability ≥ 0.95. We regard this support as being fairly strong and we show now in Figure 2 all four support values. High support values for the same clade were also reported by Rawlinson et al., 2019, and Ramirez et al., 2016. The latter regard cnidops even as being part of xenopsins. Nonetheless, we are interpreting this result carefully since the basal branching pattern of metazoan opsins differs considerably between recent studies on opsin phylogeny. Though opsin subgroups such as ciliary opsins, xenopsins, r-opsins, tetraopsins, etc. are confirmed by many studies, the interrelationships obviously depend very much on taxon sampling, choice of outgroup and tree inference algorithms used. Unfortunately, gene structure data cannot solve the question, whether cnidops and xenopsins are sister groups, since cnidops do not contain introns. Thus, we prefer to apply the term xenopsin only for protostome sequences as we did already in Vöcking et al., 2017, and as it was done by Rawlinson et al., 2019. Irrespective of these considerations, we interpret our data and published opsin phylogenies in favor for an emergence of xenopsins latest in the stem lineage of Bilateria and not lophotrochozoans. Otherwise we would expect a sister group relationship of xenopsins to another lophotrochozoan/protostome-specific opsin clade or a clade only composed of deuterostome opsins. This is not supported by our tree nor by our gene structure data and we are not aware that any publication provides respective evidence. In difference, all other protostome opsin sequences known form part of well-supported groups like r-opsin, tetraopsins, ciliary opsins and peropsins etc. In our view the simplest explanation of published and our gene tree and gene structure data is to assume a secondary loss of xenopsins in the lineage leading to deuterostomes.

We agree to the reviewers that the topology of the xenopsin subtree suggests an early divergence into two groups. This has been suggested also by Vöcking et al., 2017, and Rawlinson et al., 2019, though in both cases with moderate or low support. We did not follow this up in the first version of the manuscript since the support for the diversification of xenopsins into the two clades xenopsin A and xenopsin B was again not high. But we agree with the reviewer that this question is relevant for the manuscript. Accordingly, we did further analyses and discuss this aspect in more detail now. The revised analysis with the increased taxon-sampling did not change the picture. Support for the split is still lower as for instance for xenopsin as a whole or any other large opsin group. Thus, we run in addition trees of 1) xenopsins only (unrooted) and trees with 2) few cililary opsins, 3) few cnidops and 4) few ciliary opsins and cnidops as outgroup to investigate how robust the clades xenopsin A and xenopsin B against changes in the outgroup. The split between xenopsin A and xenopsin B is supported in the unrooted tree and in the tree rooted with cnidops, but xenopsin B has low support in the tree rooted with cnidops and ciliary opsins and is paraphyletic in the tree rooted only with ciliary opsins. We are discussing these findings in the revised version of the manuscript (subsection “Molecular phylogeny of animal xenopsins and c-opsins”) stating that our data suggest an early diversification of xenopsins is likely, but with moderate support only.

The xenopsin sequence of M. fuliginosus groups with xenopsin B representatives in all trees. We now mention this clearly in the manuscript (subsection “Molecular phylogeny of animal xenopsins and c-opsins”) and M. fuliginosus xenopsin may indeed be the first known xenopsin B of an annelid and the first xenopsin B, for which cellular expression data exist. In addition we compare sequence motifs which are important for G-protein coupling between M. fuliginosus xenopsin B and several flatworm xenopsin B, which may not be capable to induce G-protein signaling.

Since data on protein localization exist only from members of xenopsin A, we discuss considerations on the possible targeting of M. fuliginosus xenopsin more carefully now (subsection “Evolution of bilaterian eye PRCs”). Nevertheless, the closest related opsins of M. fuliginosus xenopsin for which data on subcellular localization exist, are still those, which enter cilia.

For Malacoceros xenopsin, the reasoning that xenopsin locates to cilia does not work out because the two rhabdomeric photoreceptors that strongly express it do not have cilia (only basal bodies). Where would the opsin go? There is no evidence for 'rudimentary cilia', they are simply absent (no acetylated tubulin staining). This should be stated as is.

Basal bodies close to the apical surface of PRCs are in the literature often interpreted as remnants of formerly well-developed cilia and the term rudimentary cilia is common in this context. We agree that the term is suggestive and we avoid it now. We describe the ultrastructure now more precisely (subsection “Xenopsin is coexpressed with r-opsin in cerebral eye PRCs in larval M. fuliginosus”, Figure 6—figure supplement 1) and we discuss this aspect now in more detail (subsection “Evolution of bilaterian eye PRCs”). We leave it open, where the xenopsin is going in the eye PRCs of M. fuliginosus. It may enter cilia in those cells, which have a prominent cilium and remain in the apical area of the plasma membrane in cells with reduced cilia or it may even enter microvilli. In the latter case xenopsin would be the first known opsin group, which is capable to enter both cilia and microvilli. However, all existing data on subcellular localization of xenopsin protein so far only show that xenopsin is able to enter cilia.

The figures, supplementary figures and figure legends seem to have been mixed in the new version as compared to the previous version. The numbers had disappeared. This confusing mess must be sorted out.

We are very sorry for the inconvenience and sorted the figures appropriately.

https://doi.org/10.7554/eLife.55193.sa2

Article and author information

Author details

  1. Clemens Christoph Döring

    Sars International Centre for Marine Molecular Biology, University of Bergen, Bergen, Norway
    Contribution
    Data curation, Formal analysis, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    Contributed equally with
    Suman Kumar
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-3545-7882
  2. Suman Kumar

    Sars International Centre for Marine Molecular Biology, University of Bergen, Bergen, Norway
    Contribution
    Data curation, Formal analysis, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    Contributed equally with
    Clemens Christoph Döring
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-2280-9113
  3. Sharat Chandra Tumu

    Sars International Centre for Marine Molecular Biology, University of Bergen, Bergen, Norway
    Contribution
    Formal analysis, Investigation, Methodology, Writing - original draft
    Competing interests
    No competing interests declared
  4. Ioannis Kourtesis

    Sars International Centre for Marine Molecular Biology, University of Bergen, Bergen, Norway
    Contribution
    Formal analysis, Investigation, Methodology
    Competing interests
    No competing interests declared
  5. Harald Hausen

    Sars International Centre for Marine Molecular Biology, University of Bergen, Bergen, Norway
    Contribution
    Conceptualization, Resources, Data curation, Formal analysis, Supervision, Funding acquisition, 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
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-2788-2835

Funding

European Commission (FP7-PEOPLE-2012-ITN 317172 (NEPTUNE))

  • Harald Hausen

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We are very thankful to Yannick Schwab and Rachel Mellwig at the Electron Microscopy core facility at EMBL Heidelberg for their assistance and advice in cryo-fixation of EM samples and the Arendt lab at EMBL Heidelberg for hosting cultures of M. fuliginosus and T. inopinata in their animal facility for EM fixation.

Senior Editor

  1. Diethard Tautz, Max-Planck Institute for Evolutionary Biology, Germany

Reviewing Editor

  1. Dan Larhammar, Uppsala University, Sweden

Publication history

  1. Received: January 15, 2020
  2. Accepted: September 1, 2020
  3. Accepted Manuscript published: September 3, 2020 (version 1)
  4. Version of Record published: October 1, 2020 (version 2)

Copyright

© 2020, Döring et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

Metrics

  • 1,049
    Page views
  • 193
    Downloads
  • 1
    Citations

Article citation count generated by polling the highest count across the following sources: Crossref, PubMed Central, Scopus.

Download links

A two-part list of links to download the article, or parts of the article, in various formats.

Downloads (link to download the article as PDF)

Download citations (links to download the citations from this article in formats compatible with various reference manager tools)

Open citations (links to open the citations from this article in various online reference manager services)

  1. Further reading

Further reading

    1. Evolutionary Biology
    Ben J Hightower et al.
    Research Article Updated

    How hummingbirds hum is not fully understood, but its biophysical origin is encoded in the acoustic nearfield. Hence, we studied six freely hovering Anna’s hummingbirds, performing acoustic nearfield holography using a 2176 microphone array in vivo, while also directly measuring the 3D aerodynamic forces using a new aerodynamic force platform. We corroborate the acoustic measurements by developing an idealized acoustic model that integrates the aerodynamic forces with wing kinematics, which shows how the timbre of the hummingbird’s hum arises from the oscillating lift and drag forces on each wing. Comparing birds and insects, we find that the characteristic humming timbre and radiated power of their flapping wings originates from the higher harmonics in the aerodynamic forces that support their bodyweight. Our model analysis across insects and birds shows that allometric deviation makes larger birds quieter and elongated flies louder, while also clarifying complex bioacoustic behavior.

    1. Evolutionary Biology
    Robert Niese
    Insight

    The sounds of flying animals, such as the hum of a hummingbird as it hovers, are influenced by the unique forces generated by the flapping of their wings.