Co-option of the limb patterning program in cephalopod eye development
BMC Biology volume 20, Article number: 1 (2022)
Across the Metazoa, similar genetic programs are found in the development of analogous, independently evolved, morphological features. The functional significance of this reuse and the underlying mechanisms of co-option remain unclear. Cephalopods have evolved a highly acute visual system with a cup-shaped retina and a novel refractive lens in the anterior, important for a number of sophisticated behaviors including predation, mating, and camouflage. Almost nothing is known about the molecular-genetics of lens development in the cephalopod.
Here we identify the co-option of the canonical bilaterian limb patterning program during cephalopod lens development, a functionally unrelated structure. We show radial expression of transcription factors SP6-9/sp1, Dlx/dll, Pbx/exd, Meis/hth, and a Prdl homolog in the squid Doryteuthis pealeii, similar to expression required in Drosophila limb development. We assess the role of Wnt signaling in the cephalopod lens, a positive regulator in the developing Drosophila limb, and find the regulatory relationship reversed, with ectopic Wnt signaling leading to lens loss.
This regulatory divergence suggests that duplication of SP6-9 in cephalopods may mediate the co-option of the limb patterning program. Thus, our study suggests that this program could perform a more universal developmental function in radial patterning and highlights how canonical genetic programs are repurposed in novel structures.
In the Metazoa, homologous networks of transcription factors are necessary for the development of some analogous structures in distantly related taxa. The limb patterning program is an example of this developmental process homology [1,2,3]. The limb program was first identified in the development of the proximal-distal axis of the Drosophila leg. The transcription factor SP6-9/sp1 is upstream of other program members, Dlx/dll, Pbx/exd, Meis/hth, Dac, and Arx/ar, each required for patterning specific regions of limb outgrowth [3,4,5,6,7,8,9]. This network is necessary in both vertebrate and cephalopod limb development and is expressed in a similar proximodistal pattern in a diversity of outgrowths [1, 3, 5, 10,11,12,13,14,15,16,17,18,19,20,21,22,23,24,25]. This suggests that, although each appendage is not homologous, an outgrowth program may have been present in the ancestor. Current fossil evidence and the prevalence of limbless taxa do not support an ancestor with appendages and therefore the network’s ancestral function remains unclear [1,2,3]. Many alternative hypotheses have been proposed, including an ancestral role in the nervous system, body axis formation, and radial patterning [2, 3, 26,27,28,29,30]. To understand the nature of this homology and how these co-option events occur, experiments with better sampling across the phylogeny of animals and greater diversity of developmental context are required.
Recent work identified a duplication of SP6-9 in cephalopods . Both paralogs are expressed in the developing limb in the squid Doryteuthis pealeii, while one paralog, DpSP6-9a, shows unique expression in the lens-making cells during eye development . With SP6-9 a known regulator in the limb patterning program, this new domain of expression could result in the co-option of the program in the cephalopod eye, providing a useful heterologous developmental context to better understand the network’s function.
The image-forming eye is a classic example of biological complexity and the lens is a requisite innovation in all high-resolution visual systems [32,33,34,35,36,37,38]. Cephalopods have a single-chambered eye, morphologically convergent with the vertebrate eye, composed of a cup-shaped retina and a single refractive lens . Here we perform the first in-depth molecular description of lens development in the squid Doryteuthis pealeii, we identify spatiotemporal expression of the limb patterning program in the developing eye and lens, and we demonstrate a negative regulatory role of canonical Wnt signaling upstream of the program.
Results and discussion
Cephalopod lentigenic cell differentiation and early anterior segment heterogeneity
The anterior of the cephalopod eye, or the anterior segment, is composed primarily of lens generating cells (lentigenic cells) [40,41,42]. Lentigenic cells are arranged circumferentially around the developing lens and extend long cellular processes, fusing into plates to form the lens (Fig. 1A) [40, 41, 43,44,45]. We identified the first evidence of differentiated lentigenic cells starting at late stage 21, using a previously described nuclear morphology, unique to one of the three lentigenic cell types (LC2) (Figs. 1B and 2A) [43, 44, 46]. The number of LC2 cells continues to grow until reaching pre-hatching stage (stage 29). We performed staged in situ hybridization for a homolog of DpS-Crystallin, the most abundant family of proteins in the cephalopod lens [47, 48] (Supplemental Figure 1). The first evidence of expression correlates with changes in nuclear morphology at stage 21 (Fig. 1C).
We sought to understand the molecular heterogeneity of cells in the early developing anterior segment, of which nothing is currently known. Using previously published candidates and RNA-seq data, we performed in situ hybridization screens at stage 23 to identify unique cell populations [46, 50]. We find DpSix3/6 at stage 23 expressed in the anterior segment in the distal cells that make a central cup (cc), as well as a marginal population of cells in the most proximal tissue (pm) (Fig. 2B”, Supplemental Figure 2, Supplemental Figure 3). The proximal central cells lacking DpSix3/6 expression correspond to the LC2 population (Fig. 2A”, B”). Asymmetry along the animal anterior-posterior axis in the eye is also apparent, with enrichment on the anterior side of the animal (Fig. 2B”). We also find the gene DpLhx1/5, expressed in a distal-marginal population of cells in the anterior segment (dm), and excluded from the distal central cup cells (cc) (Fig. 2C”, Supplemental Figure 2, Supplemental Figure 3). Together these genes show distinct populations of cells present early in development and provide a helpful molecular map of the anterior segment tissue at this time point: central cup cells (cc), LC2 cells (lc2), proximal-marginal cells (pm), and distal-marginal cells (dm) (Fig. 2).
Proximal-distal limb patterning genes in the anterior segment of the cephalopod
To assess whether genes involved in appendage patterning may be required for cephalopod lens development, we identified and performed in situ hybridization for the genes Dlx, Meis, Pbx, and Dac at stages 21 and 23 (Fig. 2, Supplemental Figure 2, Supplemental Figure 3). All genes were clearly expressed in the developing anterior segment and lentigenic cells with the exception of DpDac (Fig. 2E–G, Supplemental Figure 2C-2J’, Supplemental Figure 3). We find DpSP6-9a and DpDlx have overlapping expression, in the central cup cells (cc) and all proximal cells (LC2 and pm) (Fig. 2D–E”, Supplemental Figure 3). DpMeis and DpPbx are both broadly expressed in the anterior segment during lens development, with DpPbx excluded from the LC2 cells (Fig. 2F”, G”, Supplemental Figure 3).
It is known that the transcription factor aristaless is necessary for the most distal tip of the Drosophila limb in the limb program . The evolutionary relationship of Prd-like homologs (Arx/Aristaless, Alx/Aristaless-like, Rx/Retinal Homeobox, and Hbn/Homeobrain) is ambiguous across species . We identified three candidate Prd-like genes in D. pealeii and performed in situ hybridization for all three homologs, DpHbn, DpPrdl-1, and DpPrdl-2 (Supplemental Figure 2K, L) . DpHbn is expressed in the anterior segment in the distal central cup cells (cc) while DpPrdl-1 and DpPrdl-2 are excluded from the eye (Fig. 2H” and Supplemental Figure 2C, C’, K and L, Supplemental Figure 3). DpHbn’s central, distal expression recapitulates aristaless expression in the developing Drosophila limb.
Our data show that the majority of the proximodistal patterning genes in the developing limb, including SP6-9, Dlx, Meis, Pbx, as well as the Prd-like homolog, Hbn, show expression in concentric and overlapping cell populations surrounding the developing lens in the squid (Fig. 2). This pattern of expression is similar to the bullseye-like pattern of expression of these genes in the developing Drosophila limb imaginal disc and suggests a co-option of this regulatory program for a new function: patterning the cephalopod anterior segment and lens .
Canonical Wnt signaling genes expressed during anterior segment development
The duplication of SP6-9 in cephalopods may provide a substrate for the evolution of cis-regulation, resulting in novel expression of the limb patterning program in the cephalopod lens. In Drosophila appendage outgrowth, active Wnt signaling is upstream of the expression of SP6-9 [52, 53]. To assess whether Wnt may be acting upstream in the cephalopod anterior segment or whether novel regulatory mechanisms may be at play, we performed in situ hybridization for members of the Wnt signaling pathway at stage 21 and stage 23 (Fig. 3, Supplemental Figure 4). We were interested in identifying cells in the anterior segment or in adjacent tissue that may be a source of the Wnt morphogen. We performed in situ hybridization for seven Wnt homologs, with most Wnt genes expressed in the retina (Fig. 3A’, C’, and D–G). DpWnt8, DpWnt11, and DpProtostome-specific Wnt show the most robust retinal expression (Fig. 3A’, F, and G), and DpWnt7 is the only Wnt expressed in the anterior segment (Fig. 3C). DpWnt6 showed no evidence of expression in the developing eye (data not shown). These data support the hypothesis that Wnt signals emanating from the anterior segment or neighboring tissues could regulate anterior segment development.
To identify cells with potential active Wnt signaling, we analyzed the expression of Fz genes, which encode a family of Wnt receptors. We find that DpFz receptors are expressed broadly throughout the embryo. A subset of these (e.g. DpFz1/2/7, DpFz4, and DpFz5/8) are expressed in a subset of cells in the anterior segment, while others, like DpFz9/10, are excluded from the anterior segment (Fig. 3H–K, Supplemental Figure 4). On close examination, we find that DpFz5/8 is excluded asymmetrically in the anterior segment and may be important for anterior-posterior patterning (Fig. 3J’, J”, Supplemental Figure 4D). DpFz1/2/7 is excluded from the distal-marginal cells and central cup cells and interestingly, the central cup cells lacking DpFz1/2/7 are those that express all the limb patterning program genes (Fig. 3K’, K”, Supplemental Figure 4D). These data suggested active Wnt signaling may be important in the cephalopod anterior segment.
Ectopic Wnt activation leads to the loss of the lens
To assess the hypothesis that Wnt signaling is playing a regulatory role in anterior segment development, we utilized well-characterized pharmacological compounds that act as agonists and antagonists of the Wnt pathway [54,55,56,57]. We empirically determined a working concentration of LiCl (0.15 M), CHIR99021 (250 μm), and Quercetin (50 μM). We bathed embryos in the compound or vehicle control for 24 h at stage 21, the onset of lentigenic cell differentiation, and immediately fixed thereafter. Embryos were sectioned and assessed for phenotypes. Stage 21 control embryos show a thickened anterior segment, identifiable lentigenic cells, and small lens primordia (Fig. 3L). LiCl-treated stage 21 embryos show a complete absence of lens formation: no anterior segment thickening, lentigenic cells, or lens tissue. These data suggest that ectopic Wnt pathway activation inhibits lens and anterior segment development (Fig. 3L’, Supplemental Figure 5A). CHIR99021 treatment showed similar phenotypes (Supplemental Figure 5A). We assessed LiCl treated and control animals for cell death and find little difference between control and treated eyes suggesting that toxicity is unlikely the reason for these phenotypic changes (Supplemental Figure 5B). Wnt antagonist treatments (Quercetin) starting at stage 21 show lens development unaffected (Supplemental Figure 5C).
We were interested in the consequence of activating or inhibiting the Wnt pathway on lens development after the beginning of lentigenic cell differentiation. We performed the same 24-h LiCl exposure at stage 23 and find the lens smaller and the anterior segment less thick than control animals, but lentigenic cells and lens tissue remain identifiable. This suggests that ectopic Wnt signaling does not impact cell identity in differentiated lentigenic cells (Fig. 3M, M’). In Quercetin-treated animals starting at stage 23, the anterior segment shows minor organizational defects, but lens development appears unaffected (Supplemental Figure 5C).
The lack of lens growth in stage 21 treated animals may be a result of an imposed delay in lens formation or it may be a result of the loss of lens potential. To differentiate between these possibilities we allowed treated animals to recover. We bathed experimental and control embryos, at both stages 21 and 23, for 24 h, washed out the solution, and allowed animals to develop for an additional 48 h. LiCl-treated stage 21 embryos never recover a lens (Fig. 3N, 3N’) while LiCl treated stage 23 embryos do form a small but morphologically abnormal lens (Fig. 3O, O’). This abnormal lens is larger than the lens found in animals immediately fixed after treatment, suggesting that existing lentigenic cells at stage 23 continue to contribute to lens formation and growth. However, because the stage 23 treated lens is markedly smaller than the control, it suggests that further lentigenic cell differentiation is lost in treated animals. These data suggest that ectopic Wnt signaling leads to the disruption of lens potential and the lack of proper lentigenic cell differentiation.
Despite the remarkable loss of the lens as a consequence of ectopic Wnt signaling, these data do not clearly distinguish between the loss of lentigenic cell fate or proper cell function, such as the growth of the cellular processes that form the lens. To assess if lentigenic cell fate is lost, we performed in situ hybridization experiments for DpS-Crystallin on LiCl-treated animals. We saw two types of expression phenotypes, either a significant decrease (type I) or a complete loss (type II) in DpS-Crystallin expression as compared to control (Fig. 4P, P’, and P”, Supplemental Figure 6). We find all DpS-Crystallin expression exclusively dorsal to the site of lens formation suggesting that these cells may differentiate first. These data show that ectopic Wnt signaling results in the loss of lentigenic cell fate and that our treatment may have interrupted a dorsal-to-ventral wave of differentiation in some embryos (Fig. 4A). In addition, we assessed other anterior segment markers, including DpSix3/6 and DpLhx1/5, and these genes show a consistent loss of expression in the most severe phenotypes, (Supplemental Figure 6A-6C).
Limb patterning program regulatory evolution
To address if Wnt signaling is upstream of the limb patterning program, we performed in situ hybridization of limb transcription factors after LiCl treatment (Fig. 3Q–S, Supplementary Figure 6A-6C). Similar to DpS-Crystallin expression, we again see a mild reduction (Type I) or loss and severe reduction (Type II) in region of expression. Our milder phenotypes, again, show a dorsal asymmetry, which can be most easily seen in DpSP6-9A, DpDlx, and DpHbn (Fig. 3Q, Q’, Q”; R, R’, R”; and S, S’, S”). Changes are also visible but less obvious in DpPbx and DpMeis expression, with DpPbx only showing a mild phenotype (Supplemental Figure 6A-6C). These data are consistent with the placement of Wnt signaling upstream of the limb patterning program in a negative regulatory role.
Our findings indicate that the limb patterning program has been co-opted for the anterior segment and lens development in cephalopods and that this co-option does not have a homologous upstream regulatory relationship with Wnt signaling as found in the limb [24, 53]. This change in signaling and the known duplication of SP6-9 and the novel expression of the SP6-9a paralog in the anterior segment suggests that this duplication may be a mediator of limb patterning program co-option in the anterior segment. In vertebrates, although the limb patterning transcription factors are expressed during central nervous system development, including in the vertebrate retina, they do not have a role in lens development. Our gene expression data also suggest a role for the limb patterning program in the cephalopod nervous system, including the retina. It is known that SP6-9 and Dlx are required for proper regeneration of the lens-less eye in the Planarian Schmidtea mediterranea, supporting an ancestral role in the Lophotrochozoa for these genes in eye formation . The co-option of this network in the cephalopod lens may suggest an elaboration of the ancestral nervous system or retinal tissue . This is also supported by lineage tracing data, where, early in squid development, anterior segment tissue is derived from epithelial cells contiguous with the neighboring retinal primordia . In the vertebrate case, cranial ectodermal placodes are the developmental origin of the lens . The vertebrate retina is derived from evagination of forebrain neurectoderm making it unlikely that the lens evolved as an elaboration of retinal tissue . Together this suggests that the convergent evolution of complex phenotypes relies on a diversity of developmental origins.
Finally, with little similarity between limb and lens, our work suggests that the function of the limb patterning program in a limbless ancestor may have been a more generic developmental function than outgrowth. Considering present findings, previous work, and hypotheses, we conclude that the ability to pattern in a radial fashion, as previously proposed, is a more inclusive and likely ancestral function (Fig. 4B) [2, 30]. This work shows the cephalopod lens to be a unique context for future investigation of comparative regulatory changes responsible for co-option, and for identifying the regulatory mechanisms responsible for the emergent radial pattern found in embryos across species.
Doryteuthis pealeii egg sacks were obtained from the Marine Biological Labs during the summer breading season. Egg sacks were kept in well-aerated 20-gallon aquarium tanks of artificial seawater at 32–35 ppt at a pH of 8 at 20 °C. To maintain proper aeration in tanks, some embryo egg masses were kept in mesh baskets accompanied with an aeration stone. Although not required, European guidelines for cephalopod research were followed.
Histology and TUNEL staining
Embryos were fixed at 4 °C overnight in 4% paraformaldehyde (PFA) in filter-sterilized seawater. After fixation embryos were transitioned into 15% and 30% sucrose and embedded in Tissue Freezing Medium and stored at − 80 °C. Embryos were cryosectioned in 12-μm sections, stained with Sytox-Green 1:1000 and Phalloidin 555 1:300 in PBS overnight (Molecular Probes). TUNEL stained tissue was processed after sectioning using the Click-iT TUNEL Alexa Fluor 488 kit according to the manufacturer’s instructions (Invitrogen). Embryos were mounted in VECTASHIELD Hardset antifade mounting medium and imaged on a Zeiss 880 confocal.
Homolog Identification and Phylogenetics
Genes were preliminarily identified using reciprocal BLAST with Mus musculus and Drosophila melanogaster sequences as bait with the exception of S-Crystallin where previous Doryteuthis opalescens sequences were also used . Top hits in the D. pealeii transcriptome were trimmed for coding sequence and translated to amino acid sequences. To find related sequences, BLASTp was used, searching only the RefSeq protein database in NCBI filtered for vertebrate and arthropod models, as well as spiralian models when published annotated sequences could be found. The top hits of each gene name were downloaded and aligned with our D. pealeii sequences for each tree using MAFFT in Geneious . To check sequence redundancy and proper outgroups quick trees were made using FastTree. We constructed maximum-likelihood trees on the FASRC Cannon cluster supported by the FAS Division of Science Research Computing Group at Harvard University . Using PTHREADS RAxML v.8.2.10, we ran the option for rapid bootstrapping with search for best maximum likelihood tree, resampling with 1000 bootstrap replicates, the PROTGAMMAAUTO model of amino acid substitution, and otherwise default parameters . Fasta alignments, Nexus tree files are found at doi:10.5061/dryad.vhhmgqnvf. All PDF versions of the trees are found in Supplemental Figure 1.
Cloning and probe synthesis
Embryos stage 21–29 were crushed in TRIzol reagent. RNA was extracted using standard phenol-chloroform extraction with a clean-up using the Qiagen RNeasy Micro kit. cDNA was synthesized using iScript (Bio-Rad) according to manufacturer protocols. Primers were designed using Primer3 in the Geneious software package from available transcriptomic data (Koenig et al., 2016). PCR products were ligated into the Pgem-T Easy plasmid and isolated using the Qiagen miniprep kit. Plasmids were linearized using restriction enzymes. Sense and anti-sense probes were synthesized using T7 and SP6 polymerase with digoxygenin-labeled nucleotides.
In situ hybridization
Embryos were fixed as previously described  and were dehydrated in 100% ethanol and stored at − 20 °C. Whole-mount in situ hybridization was performed as previously described . Embryos were imaged using a Zeiss Axio Zoom.V16. Embryos were fixed for sectioning overnight in 4% PFA in artificial seawater and dehydrated in 100% ethanol. Embryos were transitioned into histoclear and embedded in paraffin. Embryos were sectioned on a Leica RM2235 microtome in 5-μm sections. Sections were dewaxed for in situ in Histoclear, rehydrated through an EtOH series, and re-fixed for 5 min at 4 °C in 4% PFA in PBS. Embryos were exposed to Proteinase K for 20 min at 37 °C and then quenched with glycine. The embryos were then de-acetylated with acetic anhydride. Slides were then pre-hybridized at 65 °C for 30–60 min and then exposed to probe overnight. Slides were washed in 50% formamide/1× SSC/0.1% Tween-20 hybridization buffer twice, then twice in 1× SSC, .2× SSC, and 0.02× SSC, all at 70 °C. The slides were then washed at room temperature in MABT three times and blocked in Roche Blocking Buffer for an hour. Slides were incubated in Anti-Dig antibody (Roche) at 1/4000 overnight at 4 °C. Slides were washed with MABT and then placed in AP reaction buffer. Slides were then exposed to BCIP/NBT solution until reacted and stopped in PBS. Slides were counterstained with Sytox-Green 1:1000 overnight. Slides mounted in ImmunoHistoMount (Abcam) and imaged on a Zeiss Axioscope. DpS-Crystallin embryo in situs were transitioned to sucrose and embedded after imaging in whole-mount. Embryos were image on a Zeiss Axioscope.
Ex ovo experimental culture
Ex ovo culture was performed as previously described . Embryos were bathed in .25 M, .15 M, and .07 M LiCl; 100 nm, 250 nm, and 500 nm concentration of Wnt Agonist (CHIR99021); and 25 μM, 50 μM, and 100 μM Quercetin in Pen-Step filter-sterilized seawater to determine a working concentration. Control animals were bathed in equivalent amounts of DMSO or Pen-Strep alone.
Availability of data and materials
All sequences generated and analyzed in this study have been deposited in NCBI’s GenBank database under accession numbers MZ020516-MZ020549. All multiple sequence alignments and phylogenetic trees are available at doi:10.5061/dryad.vhhmgqnvf .
- a :
- aco :
Anterior chamber organ
- as :
- cc :
- dm :
- e :
- f :
- l :
- lc2 :
LC2 lentigenic cell population
- lp :
- m :
- mo :
- pm :
- rp :
- y :
Shubin N, Tabin C, Carroll S. Fossils, genes and the evolution of animal limbs. Nature. 1997;388(6643):639–48. https://doi.org/10.1038/41710.
Erwin DH, Davidson EH. The last common bilaterian ancestor. Development. 2002;129:3021–32 https://www.ncbi.nlm.nih.gov/pubmed/12070079.
Pueyo JI, Couso JP. Parallels between the proximal-distal development of vertebrate and arthropod appendages: homology without an ancestor? Curr Opin Genet Dev. 2005;15(4):439–46. https://doi.org/10.1016/j.gde.2005.06.007.
Panganiban G, Nagy L, Carroll SB. The role of the Distal-less gene in the development and evolution of insect limbs. Curr Biol. 1994;4(8):671–5. https://doi.org/10.1016/s0960-9822(00)00151-2.
Panganiban G, Irvine SM, Lowe C, Roehl H, Corley LS, Sherbon B, et al. The origin and evolution of animal appendages. Proceedings of the National Academy of Sciences. 1997;94(10):5162–6. https://doi.org/10.1073/pnas.94.10.5162.
Dong PD, Chu J, Panganiban G. Proximodistal domain specification and interactions in developing Drosophila appendages. Development. 2001;128(12):2365–72. https://www.ncbi.nlm.nih.gov/pubmed/11493555. https://doi.org/10.1242/dev.128.12.2365.
Dong PDS, Dicks JS, Panganiban G. Distal-less and homothorax regulate multiple targets to pattern the Drosophila antenna. Development. 2002;129(8):1967–74. https://www.ncbi.nlm.nih.gov/pubmed/11934862. https://doi.org/10.1242/dev.129.8.1967.
Estella C, Voutev R, Mann RS. A dynamic network of morphogens and transcription factors patterns the fly leg. Curr Top Dev Biol. 2012;98:173–198. doi:https://doi.org/10.1016/B978-0-12-386499-4.00007-0.
Campbell G, Tomlinson A. The roles of the homeobox genes aristaless and Distal-less in patterning the legs and wings of Drosophila. Development. 1998;125(22):4483–93. https://www.ncbi.nlm.nih.gov/pubmed/9778507. https://doi.org/10.1242/dev.125.22.4483.
Maas R, Bei M. The Genetic Control of Early Tooth Development. Critical Reviews in Oral Biology & Medicine. 1997;8(1):4–39. https://doi.org/10.1177/10454411970080010101.
Mercader N, Leonardo E, Azpiazu N, Serrano A, Morata G, Martínez C, et al. Conserved regulation of proximodistal limb axis development by Meis1/Hth. Nature. 1999;402(6760):425–9. https://doi.org/10.1038/46580.
Panganiban G, Rubenstein JLR. Developmental functions of the Distal-less/Dlx homeobox genes. Development. 2002;129(19):4371–86. https://www.ncbi.nlm.nih.gov/pubmed/12223397. https://doi.org/10.1242/dev.129.19.4371.
Prpic N-M, Tautz D. The expression of the proximodistal axis patterning genes Distal-less and dachshund in the appendages of Glomeris marginata (Myriapoda: Diplopoda) suggests a special role of these genes in patterning the head appendages. Dev Biol. 2003;260(1):97–112. https://doi.org/10.1016/s0012-1606(03)00217-3.
Angelini DR, Kaufman TC. Insect appendages and comparative ontogenetics. Dev Biol. 2005;286(1):57–77. https://doi.org/10.1016/j.ydbio.2005.07.006.
Shubin N, Tabin C, Carroll S. Deep homology and the origins of evolutionary novelty. Nature. 2009;457(7231):818–23. https://doi.org/10.1038/nature07891.
Moczek AP, Rose DJ. Differential recruitment of limb patterning genes during development and diversification of beetle horns. Proc Natl Acad Sci U S A. 2009;106(22):8992–7. https://doi.org/10.1073/pnas.0809668106.
Capellini TD, Zappavigna V, Selleri L. Pbx homeodomain proteins: TALEnted regulators of limb patterning and outgrowth. Dev Dyn. 2011;240(5):1063–86. https://doi.org/10.1002/dvdy.22605.
Ibarretxe G, Aurrekoetxea M, Crende O, Badiola I, Jimenez-Rojo L, Nakamura T, et al. Epiprofin/Sp6 regulates Wnt-BMP signaling and the establishment of cellular junctions during the bell stage of tooth development. Cell Tissue Res. 2012;350(1):95–107. https://doi.org/10.1007/s00441-012-1459-8.
Grimmel J, Dorresteijn AWC, Fröbius AC. Formation of body appendages during caudal regeneration in Platynereis dumerilii: adaptation of conserved molecular toolsets. Evodevo. 2016;7(1):10. https://doi.org/10.1186/s13227-016-0046-6.
Sanz-Navarro M, Delgado I, Torres M, Mustonen T, Michon F, Rice DP. Dental Epithelial Stem Cells Express the Developmental Regulator Meis1. Frontiers in Physiology. 2019;10. doi:https://doi.org/10.3389/fphys.2019.00249.
Ramanathan A, Srijaya TC, Sukumaran P, Zain RB, Kasim NHA. Homeobox genes and tooth development: Understanding the biological pathways and applications in regenerative dental science. Archives of Oral Biology. 2018;85:23–39. doi:https://doi.org/10.1016/j.archoralbio.2017.09.033.
Setton EVW, Sharma PP. Cooption of an appendage-patterning gene cassette in the head segmentation of arachnids. Proc Natl Acad Sci U S A. 2018;115(15):E3491–500. https://doi.org/10.1073/pnas.1720193115.
Minelli A. Limbs and tail as evolutionarily diverging duplicates of the main body axis. Evol Dev. 2000;2(3):157–65. https://doi.org/10.1046/j.1525-142x.2000.00054.x.
Lemons D, Fritzenwanker JH, Gerhart J, Lowe CJ, McGinnis W. Co-option of an anteroposterior head axis patterning system for proximodistal patterning of appendages in early bilaterian evolution. Dev Biol. 2010;344(1):358–62. https://doi.org/10.1016/j.ydbio.2010.04.022.
McDougall C, Korchagina N, Tobin JL, Ferrier DE. Annelid Distal-less/Dlx duplications reveal varied post-duplication fates. BMC Evol Biol. 2011;11(1):241. https://doi.org/10.1186/1471-2148-11-241.
Plavicki JS, Squirrell JM, Eliceiri KW, Boekhoff-Falk G. Expression of the Drosophila homeobox gene, Distal-less, supports an ancestral role in neural development. Dev Dyn. 2016;245(1):87–95. https://doi.org/10.1002/dvdy.24359.
Carroll SB, Gates J, Keys DN, Paddock SW, Panganiban GE, Selegue JE, et al. Pattern formation and eyespot determination in butterfly wings. Science. 1994;265(5168):109–14. https://doi.org/10.1126/science.7912449.
McCulloch KJ, Koenig KM. Krüppel-like factor/specificity protein evolution in the Spiralia and the implications for cephalopod visual system novelties. Proc Biol Sci. 2020;287(1937):20202055. https://doi.org/10.1098/rspb.2020.2055.
Arendt D, Hausen H, Purschke G. The “division of labour” model of eye evolution. Philos Trans R Soc Lond B Biol Sci. 2009;364(1531):2809–17. https://doi.org/10.1098/rstb.2009.0104.
The eyes of pecten, spondylus, amussium and allied lamellibranchs, with a short discussion on their evolution. Proceedings of the Royal Society of London. Series B, Containing Papers of a Biological Character. 1928;103:355–65. doi:https://doi.org/10.1098/rspb.1928.0047.
Walls GL. ORIGIN OF THE VERTEBRATE EYE. Archives of Ophthalmology. 1939;22:452–486. doi:https://doi.org/10.1001/archopht.1939.00860090118018.
Koenig KM, Gross JM. Evolution and development of complex eyes: a celebration of diversity. Development. 2020;147(19). https://doi.org/10.1242/dev.182923.
Nilsson D-E. Eye evolution and its functional basis. Vis Neurosci. 2013;30(1-2):5–20. https://doi.org/10.1017/S0952523813000035.
Jonasova K, Kozmik Z. Eye evolution: Lens and cornea as an upgrade of animal visual system. Seminars in Cell & Developmental Biology. 2008;19(2):71–81. https://doi.org/10.1016/j.semcdb.2007.10.005.
Packard A. CEPHALOPODS AND FISH: THE LIMITS OF CONVERGENCE. Biological Reviews. 1972;47:241–307. doi:https://doi.org/10.1111/j.1469-185x.1972.tb00975.x.
Williams LW. The Anatomy of the common squid. Loligo Pealii: Lesueur, by Leonard Worcester Williams; 1909. https://books.google.com/books/about/The_Anatomy_of_the_common_squid_Loligo_P.html?hl=&id=rrbFPgAACAAJ. https://doi.org/10.5962/bhl.title.27291.
Arnold JM. Fine structure of the development of the cephalopod lens. J Ultrastruct Res. 1967;17(5-6):527–43. https://www.ncbi.nlm.nih.gov/pubmed/6025339. https://doi.org/10.1016/S0022-5320(67)80139-4.
Arnold JM. On the occurrence of microtubules in the developing lens of the squid Loligo pealii. J Ultrastruct Res. 1966;14(5-6):534–9. https://www.ncbi.nlm.nih.gov/pubmed/5930349. https://doi.org/10.1016/S0022-5320(66)80080-1.
West JA, Sivak JG, Doughty MJ. Microscopical evaluation of the crystalline lens of the squid (Loligo opalescens) during embryonic development. Exp Eye Res. 1995;60(1):19–35. https://doi.org/10.1016/s0014-4835(05)80080-6.
Meinertzhagen IA. Development of the Squid’s Visual System. Squid as Experimental Animals. 1990:399–419. https://doi.org/10.1007/978-1-4899-2489-6_18.
Chiou SH. Physicochemical characterization of a crystallin from the squid lens and its comparison with vertebrate lens crystallins. J Biochem. 1984;95(1):75–82. https://doi.org/10.1093/oxfordjournals.jbchem.a134605.
West JA, Sivak JG, Pasternak J, Piatigorsky J. Immunolocalization of S-crystallins in the developing squid (Loligo opalescens) lens. Dev Dyn. 1994;199(2):85–92. https://doi.org/10.1002/aja.1001990202.
Arnold JM. NORMAL EMBRYONIC STAGES OF THE SQUID, LOLIGO PEALII (LESUEUR). Biol Bull. 1965;128(1):24–32. https://doi.org/10.2307/1539386.
Ogura A, Yoshida M-A, Moritaki T, Okuda Y, Sese J, Shimizu KK, et al. Loss of the six3/6 controlling pathways might have resulted in pinhole-eye evolution in Nautilus. Sci Rep. 2013;3(1):1432. https://doi.org/10.1038/srep01432.
Schiemann SM, Martín-Durán JM, Børve A, Vellutini BC, Passamaneck YJ, Hejnol A. Clustered brachiopod Hox genes are not expressed collinearly and are associated with lophotrochozoan novelties. doi:https://doi.org/10.1101/058669.
Cohen SM. Specification of limb development in the Drosophila embryo by positional cues from segmentation genes. Nature. 1990;343(6254):173–7. https://doi.org/10.1038/343173a0.
Estella C, Rieckhof G, Calleja M, Morata G. The role of buttonhead and Sp1 in the development of the ventral imaginal discs of Drosophila. Development. 2003;130(24):5929–41. https://doi.org/10.1242/dev.00832.
Hedgepeth CM, Conrad LJ, Zhang J, Huang HC, Lee VM, Klein PS. Activation of the Wnt signaling pathway: a molecular mechanism for lithium action. Dev Biol. 1997;185(1):82–91. https://doi.org/10.1006/dbio.1997.8552.
Klein PS, Melton DA. A molecular mechanism for the effect of lithium on development. Proceedings of the National Academy of Sciences. 1996;93(16):8455–9. https://doi.org/10.1073/pnas.93.16.8455.
Sato N, Meijer L, Skaltsounis L, Greengard P, Brivanlou AH. Maintenance of pluripotency in human and mouse embryonic stem cells through activation of Wnt signaling by a pharmacological GSK-3-specific inhibitor. Nat Med. 2004;10(1):55–63. https://doi.org/10.1038/nm979.
Park CH, Chang JY, Hahm ER, Park S, Kim H-K, Yang CH. Quercetin, a potent inhibitor against β-catenin/Tcf signaling in SW480 colon cancer cells. Biochemical and Biophysical Research Communications. 2005;328(1):227–34. https://doi.org/10.1016/j.bbrc.2004.12.151.
Cvekl A, Ashery-Padan R. The cellular and molecular mechanisms of vertebrate lens development. Development. 2014;141(23):4432–47. https://doi.org/10.1242/dev.107953.
Chow RL, Lang RA. Early eye development in vertebrates. Annu Rev Cell Dev Biol. 2001;17(1):255–96. https://doi.org/10.1146/annurev.cellbio.17.1.255.
Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ. Basic local alignment search tool. J Mol Biol. 1990;215(3):403–10. https://doi.org/10.1016/S0022-2836(05)80360-2.
Katoh K, Misawa K, Kuma K-I, Miyata T. MAFFT: a novel method for rapid multiple sequence alignment based on fast Fourier transform. Nucleic Acids Res. 2002;30(14):3059–66. https://doi.org/10.1093/nar/gkf436.
Price MN, Dehal PS, Arkin AP. FastTree 2--approximately maximum-likelihood trees for large alignments. PLoS One. 2010;5(3):e9490. https://doi.org/10.1371/journal.pone.0009490.
Stamatakis A. RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics. 2014;30(9):1312–3. https://doi.org/10.1093/bioinformatics/btu033.
Koenig KM. Multiple sequence alignments and phylogenetic trees from: Co-option of the limb patterning program in cephalopod eye development, Dryad, Dataset, https://doi.org/10.5061/dryad.vhhmgqnvf 2021.
The authors would like to thank the Koenig and Srivastava lab members for helpful discussions as well as Kevin Woods and the John Harvard Distinguished Science Fellows community for support. We would like to thank Jeffrey Gross, Alex Schier, Mansi Srivastava, Nick Bellono, and Andrew Murray for comments on the manuscript. We also thank the Marine Biological Labs, the Marine Resources Center, Owen Nichols, Ernie Eldredge, and Shannon Eldredge for assisting in the acquisition of embryos. We would also like to acknowledge the Harvard College undergraduates of LS50: Integrated Science Laboratory Course: Zach Alerte, Vlad Batagui, Eli Burnes, Stephen Casper, Chris Chen, Ahab Chopra, Ralph Estanboulieh, Lily Gao, Pedro Garcia, Saimun Habib, Harry Hager, Maxwell Ho, Charlie Horowitz, Ray Jiang, Prashanth Kumar, Truelian Lee, Arian Mansur, Matthew Mardo, Mark Theodore Meneses, Kendrick Nguyen, Francesco Rolando, Simon Schnabl, Taylor Shirtliff-Hinds, Sorscher Lincoln, William Stainier, Avi Swartz, David Szanto, Sophia Tang, Joey Toker, Analli Torres, Nina Uzoigwe, Rowen VonPlagenhoef, Evelyn Wong, Alexandra Zaloga, Maxwell Zhu.
This work is supported by the Office of the NIH Director 1DP5OD023111-01, John Harvard Distinguished Science Fellowship to K.M.K.
Ethics approval and consent to participate
No ethics approvals or consent is required to work with cephalopod embryos.
Consent for publication
The authors declare that they have no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
includes supplemental Figures 1-6. Supplemental Figure 1 is Maximum-likelihood phylogenetic trees for genes identified in this study. Supplemental Figure 2 is limb network supplemental data. Supplemental Figure 3 is targeted image enlargement of anterior segment gene expression. Supplemental Figure 4 is Wnt signaling expression supplemental data. Supplemental Figure 5 is Wnt agonist and antagonist supplemental data. Supplemental Figure 6 Supplemental in situ hybridization data and quantification. Supplemental Table 1 is are the primer sequences used to clone the genes included in this study.
About this article
Cite this article
Neal, S., McCulloch, K.J., Napoli, F.R. et al. Co-option of the limb patterning program in cephalopod eye development. BMC Biol 20, 1 (2022). https://doi.org/10.1186/s12915-021-01182-2