A conserved Shh cis-regulatory module highlights a common developmental origin of unpaired and paired fins

  • Nature Geneticsvolume 50pages504509 (2018)
  • doi:10.1038/s41588-018-0080-5
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Despite their evolutionary, developmental and functional importance, the origin of vertebrate paired appendages remains uncertain. In mice, a single enhancer termed ZRS is solely responsible for Shh expression in limbs. Here, zebrafish and mouse transgenic assays trace the functional equivalence of ZRS across the gnathostome phylogeny. CRISPR/Cas9-mediated deletion of the medaka (Oryzias latipes) ZRS and enhancer assays identify the existence of ZRS shadow enhancers in both teleost and human genomes. Deletion of both ZRS and shadow ZRS abolishes shh expression and completely truncates pectoral fin formation. Strikingly, deletion of ZRS results in an almost complete ablation of the dorsal fin. This finding indicates that a ZRS-Shh regulatory module is shared by paired and median fins and that paired fins likely emerged by the co-option of developmental programs established in the median fins of stem gnathostomes. Shh function was later reinforced in pectoral fin development with the recruitment of shadow enhancers, conferring additional robustness.

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  1. 1.

    Freitas, R., Gomez-Skarmeta, J. L. & Rodrigues, P. N. New frontiers in the evolution of fin development. J. Exp. Zool. B Mol. Dev. Evol. 322, 540–552 (2014).

  2. 2.

    Sordino, P., van der Hoeven, F. & Duboule, D. Hox gene expression in teleost fins and the origin of vertebrate digits. Nature 375, 678–681 (1995).

  3. 3.

    Neumann, C. J., Grandel, H., Gaffield, W., Schulte-Merker, S. & Nüsslein-Volhard, C. Transient establishment of anteroposterior polarity in the zebrafish pectoral fin bud in the absence of sonic hedgehog activity. Development 126, 4817–4826 (1999).

  4. 4.

    Ahn, D. G., Kourakis, M. J., Rohde, L. A., Silver, L. M. & Ho, R. K. T-box genetbx5 is essential for formation of the pectoral limb bud. Nature 417, 754–758 (2002).

  5. 5.

    Dahn, R. D., Davis, M. C., Pappano, W. N. & Shubin, N. H. Sonic hedgehog function in chondrichthyan fins and the evolution of appendage patterning. Nature 445, 311–314 (2007).

  6. 6.

    Freitas, R., Zhang, G. & Cohn, M. J. Evidence that mechanisms of fin development evolved in the midline of early vertebrates. Nature 442, 1033–1037 (2006).

  7. 7.

    Pieretti, J. et al. Organogenesis in deep time: a problem in genomics, development, and paleontology. Proc. Natl. Acad. Sci. USA 112, 4871–4876 (2015).

  8. 8.

    Wittkopp, P. J. & Kalay, G. Cis-regulatory elements: molecular mechanisms and evolutionary processes underlying divergence. Nat. Rev. Genet. 13, 59–69 (2011).

  9. 9.

    Peter, I. S. & Davidson, E. H. Evolution of gene regulatory networks controlling body plan development. Cell 144, 970–985 (2011).

  10. 10.

    Gehrke, A. R. & Shubin, N. H. Cis-regulatory programs in the development and evolution of vertebrate paired appendages. Semin. Cell Dev. Biol. 57, 31–39 (2016).

  11. 11.

    Ogura, T. et al. Evidence that Shh cooperates with a retinoic acid inducible co-factor to establish ZPA-like activity. Development 122, 537–542 (1996).

  12. 12.

    Krauss, S., Concordet, J. P. & Ingham, P. W. A functionally conserved homolog of the Drosophila segment polarity gene hh is expressed in tissues with polarizing activity in zebrafish embryos. Cell 75, 1431–1444 (1993).

  13. 13.

    Lettice, L. A. et al. A long-range Shh enhancer regulates expression in the developing limb and fin and is associated with preaxial polydactyly. Hum. Mol. Genet. 12, 1725–1735 (2003).

  14. 14.

    Sagai, T., Hosoya, M., Mizushina, Y., Tamura, M. & Shiroishi, T. Elimination of a long-range cis-regulatory module causes complete loss of limb-specific Shh expression and truncation of the mouse limb. Development 132, 797–803 (2005).

  15. 15.

    Maas, S. A., Suzuki, T. & Fallon, J. F. Identification of spontaneous mutations within the long-range limb-specific Sonic hedgehog enhancer (ZRS) that alter Sonic hedgehog expression in the chicken limb mutants oligozeugodactyly and silkie breed. Dev. Dyn. 240, 1212–1222 (2011).

  16. 16.

    Chiang, C. et al. Cyclopia and defective axial patterning in mice lacking Sonic hedgehog gene function. Nature 383, 407–413 (1996).

  17. 17.

    Kvon, E. Z. et al. Progressive loss of function in a limb enhancer during snake evolution. Cell 167, 633–642.e11 (2016).

  18. 18.

    Irimia, M. et al. Comparative genomics of the Hedgehog loci in chordates and the origins of Shh regulatory novelties. Sci. Rep. 2, 433 (2012).

  19. 19.

    Jaillon, O. et al. Genome duplication in the teleost fish Tetraodon nigroviridis reveals the early vertebrate proto-karyotype. Nature 431, 946–957 (2004).

  20. 20.

    Amores, A. et al. Zebrafish hox clusters and vertebrate genome evolution. Science 282, 1711–1714 (1998).

  21. 21.

    Lettice, L. A. et al. Development of five digits is controlled by a bipartite long-range cis-regulator. Development 141, 1715–1725 (2014).

  22. 22.

    Gehrke, A. R. et al. Deep conservation of wrist and digit enhancers in fish. Proc. Natl. Acad. Sci. USA 112, 803–808 (2015).

  23. 23.

    Cotney, J. et al. The evolution of lineage-specific regulatory activities in the human embryonic limb. Cell 154, 185–196 (2013).

  24. 24.

    Westerfield, M. The Zebrafish Book (University of Oregon Press, Eugene, Oregon, USA, 1995).

  25. 25.

    Koster, R., Stick, R., Loosli, F. & Wittbrodt, J. Medaka spalt acts as a target gene of hedgehog signaling. Development 124, 3147–3156 (1997).

  26. 26.

    Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B. & Schilling, T. F. Stages of embryonic development of the zebrafish. Dev. Dyn. 203, 253–310 (1995).

  27. 27.

    Iwamatsu, T. Stages of normal development in the medaka Oryzias latipes. Mech. Dev. 121, 605–618 (2004).

  28. 28.

    Schneider, I. et al. Appendage expression driven by the Hoxd Global Control Region is an ancient gnathostome feature. Proc. Natl. Acad. Sci. USA 108, 12782–12786 (2011).

  29. 29.

    Bird, N. C. & Mabee, P. M. Developmental morphology of the axial skeleton of the zebrafish, Danio rerio (Ostariophysi: Cyprinidae). Dev. Dyn. 228, 337–357 (2003).

  30. 30.

    Acemel, R. D. et al. A single three-dimensional chromatin compartment in amphioxus indicates a stepwise evolution of vertebrate Hox bimodal regulation. Nat. Genet. 48, 336–341 (2016).

  31. 31.

    Kawakami, K. et al. A transposon-mediated gene trap approach identifies developmentally regulated genes in zebrafish. Dev. Cell 7, 133–144 (2004).

  32. 32.

    Jowett, T. & Lettice, L. Whole-mount in situ hybridizations on zebrafish embryos using a mixture of digoxigenin- and fluorescein-labelled probes. Trends Genet. 10, 73–74 (1994).

  33. 33.

    Martinez-Morales, J. R. et al. Differentiation of the vertebrate retina is coordinated by an FGF signaling center. Dev. Cell 8, 565–574 (2005).

  34. 34.

    Fernández-Miñán, A., Bessa, J., Tena, J. J. & Gómez-Skarmeta, J. L. Assay for transposase-accessible chromatin and circularized chromosome conformation capture, two methods to explore the regulatory landscapes of genes in zebrafish. Methods Cell Biol. 135, 413–430 (2016).

  35. 35.

    Moreno-Mateos, M. A. et al. CRISPRscan: designing highly efficient sgRNAs for CRISPR-Cas9 targeting in vivo. Nat. Methods 12, 982–988 (2015).

  36. 36.

    Stemmer, M. & Thumberger, T. Del Sol Keyer, M., Wittbrodt, J. & Mateo, J. L. CCTop: an intuitive, flexible and reliable CRISPR/Cas9 target prediction tool. PLoS One 10, e0124633 (2015).

  37. 37.

    Vejnar, C. E., Moreno-Mateos, M. A., Cifuentes, D., Bazzini, A. A. & Giraldez, A. J. Optimized CRISPR-Cas9 System for genome editing in zebrafish. Cold Spring Harb. Protoc. 2016, pdb.prot086850 (2016).

  38. 38.

    Gagnon, J. A. et al. Efficient mutagenesis by Cas9 protein-mediated oligonucleotide insertion and large-scale assessment of single-guide RNAs. PLoS One 9, e98186 (2014).

  39. 39.

    Sugahara, F., Murakami, Y. & Kuratani, S. Gene expression analysis of lamprey embryos. in In Situ Hybridization Methods (ed. Hauptmann, G.) 263–278 (Springer, New York, 2015).

  40. 40.

    Tahara, Y. Normal stages of development in the lamprey, Lampetra reissneri (Dybowski). Zool. Sci. 5, 109–118 (1988).

  41. 41.

    Sugahara, F. et al. Evidence from cyclostomes for complex regionalization of the ancestral vertebrate brain. Nature 531, 97–100 (2016).

  42. 42.

    Kusakabe, R., Takechi, M., Tochinai, S. & Kuratani, S. Lamprey contractile protein genes mark different populations of skeletal muscles during development. J. Exp. Zool. B Mol. Dev. Evol. 302, 121–133 (2004).

  43. 43.

    Ohtani, K. et al. Expression of Sox and fibrillar collagen genes in lamprey larval chondrogenesis with implications for the evolution of vertebrate cartilage. J. Exp. Zool. B Mol. Dev. Evol. 310, 596–607 (2008).

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We thank A. Fernández-Miñan from the CABD aquatic vertebrate platform for providing the medaka 4C-seq samples, R.D. Acemel for helping with the design of the medaka 4C-seq primers, and all members of JLGSK laboratory and F. Casares for discussions. We thank J. Westlund for illustration assistance and F. Sugahara for providing the clone of lamprey HhA and the metamorphic ammocoete larva. This project received funding from the European Research Council under the European Union’s Horizon 2020 research and innovation programme (grant agreement No. 740041), the European Union’s Horizon 2020 research and innovation programme under the Marie Sklodowska-Curie grant agreement #658521, the Spanish Ministerio de Economía y Competitividad (grants BFU2016-74961-P, BFU2014-53765-P and BFU2016-81887-REDT), the Andalusian Government (grant BIO-396), CNPq Universal Program Grant 403248/2016-7 and CAPES/Alexander von Humboldt Foundation fellowship (to I.S.). J.L. was supported by Becas Chile.

Author information

Author notes

  1. These authors contributed equally: Joaquín Letelier, Elisa de la Calle-Mustienes.


  1. Centro Andaluz de Biología del Desarrollo (CABD), Consejo Superior de Investigaciones Científicas/Universidad Pablo de Olavide/Junta de Andalucía, Sevilla, Spain

    • Joaquín Letelier
    • , Elisa de la Calle-Mustienes
    • , Silvia Naranjo
    • , Ignacio Maeso
    • , Juan Ramón Martinez-Morales
    •  & José Luis Gómez-Skarmeta
  2. Department of Organismal Biology and Anatomy, University of Chicago, Chicago, IL, USA

    • Joyce Pieretti
    • , Tetsuya Nakamura
    •  & Neil H. Shubin
  3. Evolutionary Morphology Laboratory, RIKEN, Kobe, Japan

    • Juan Pascual-Anaya
  4. Marine Biological Laboratory, Woods Hole, MA, USA

    • Neil H. Shubin
  5. Instituto de Ciências Biológicas, Universidade Federal do Pará, Belém, Brazil

    • Igor Schneider


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J.L. generated and analyzed the medaka mutants. E.d.l.C.-M. carried out the 4C-seq experiments and the zebrafish transgenic assays with the help of S.N. and J.L. J.P. generated the mouse transgenic data. T.N. performed the µCT experiments. J.P.-A. carried out the lamprey in situ experiments. J.L.G.-S., J.R.M.-M., I.S. and N.H.S. conceived, designed and coordinated the project with the help of J.L. and I.M.  I.S., J.L.G.-S., J.R.M.-M., N.H.S., J.L. and I.M. wrote the manuscript.

Competing interests

The authors declare no competing interests.

Corresponding authors

Correspondence to Neil H. Shubin or Igor Schneider or Juan Ramón Martinez-Morales or José Luis Gómez-Skarmeta.

Integrated supplementary information

  1. Supplementary Figure 1 Sequence conservation and enhancer activity of the ZRS from different species and the corresponding amphioxus Lmbr1 intron in zebrafish fins and mouse limbs.

    a, the ZRS enhancer is located between Exon 5 and 6 of the Lmbr1 gene. b, vista plot showing the conservation degree of the ZRS enhancer in different species. c-f, transgenic mouse embryos showing the activity of the ZRS region from different species in the E10.5 developing limbs. Scale bars, 200μm. g-k, stable transgenic zebrafish larvae showing the activity of the ZRS region from different species at 72hpf pectoral fins. Arrows point to the ZPA domain. Scale bars, 50μm. For each construct shown in c-k, three or more independent transgenic lines were generated.

  2. Supplementary Figure 2 Hh expression pattern in the dorsal fins of the lamprey ammocoete larva during metamorphosis.

    a, b, first dorsal (a) and second dorsal and caudal fins (b) of an ammocoete larva of the lamprey L. reissneri. Scale bars, 5 mm. c-g, in situ hybridizations of L. reissneri first (c, d) and second (e-g) dorsal fins in sections at different anterior-posterior levels (as shown in a and b), assayed with an equimolecular mix of digoxigenin-labeled riboprobes of the lamprey L. camtschaticum HhA, HhB and HhD genes. No expression of Hh genes in the underlying mesenchyme of developing dorsal fins was detected. Scale bars, 100 μm. h-j, L. camtschaticum whole embryos at stage 27 hybridized with individual riboprobes of HhA (h), HhB (i) and HhD (j) used as positive controls for the Hh genes’ probes. HhD expression pattern has not been hitherto described (see Sugahara et al., 2016). Scale bars, 200μm. k, l, section in situ hybridizations of L. reissneri second dorsal fin, at the levels shown in b, with digoxigenin-labeled riboprobes of the L. camtschaticum MyHC1 (k) and ColA (l) genes (used as positive control). Expression in the skeletal muscle (MyHC1) and cartilage (ColA), was observed. Scale bars, 200 μm. Lamprey expression experiments were repeated two times independently with similar results.

  3. Supplementary Figure 3 µCT scanning of adult pectoral fins in wild-type and ∆401 mutant fish.

    a, b the architecture of fin rays and proximal radials was revealed after scanning for three-month-old wild type and medaka mutants. Note the reduced number of endoskeletal elements in the mutants. µCT scanning experiments were performed for three WT and five ∆401 ZRS adult fish. Scale bars, 500μm c, in contrast to the endoskeletal defects, measurement of fin ray length did not show any significant differences (p-value=0.122) between wild type (mean=0.185) and ∆948 mutant fish (mean=0.204). A t-test was used for analysis of pectoral fin length measurements. Each point in the graph represent pectoral fin length measurements from independent animals.

  4. Supplementary Figure 4 Potential fin and limb enhancers in the introns of the zebrafish and human lmbr1 and LMBR1 genes.

    Genome coordinates are shown in the x axis and reads counts in the y axis. Several potential fin/limb enhancers, as predicted by H3K27ac signature are found in different introns of the zebrafish and human lmbr1/LMBR1 genes. Although conserved with other mammals, these enhancers are not conserved in the mouse genome (red arrowheads). We search for putative ETS transcription factor binding sites in the sZRS of the three different species tested in the transgenesis assay. We found 3 ETS transcription-binding sites in the sZRS from zebrafish, 7 sites in medaka sZRS and 9 in human sZRS.

  5. Supplementary Figure 5 Progression of pectoral fin development is truncated in ∆3.4-kb ZRS mutants.

    a-c, temporal series showing normal development of pectoral fins in wild type embryos (black arrowheads). d-f, at 3dpf pectoral fin buds appears reduced (black arrowhead) in mutant embryos and are absent during later developmental stages (asterisks). We could not analyse in detail the mature histology of other fins in the double mutant (ZRS-sZRS) due to fish lethality before the onset of pelvic and anal fins maturation (from 3–4 weeks post fertilization on). Nevertheless, the lack of activity of the sZRS in the anal fin (Extended data Figure 7d) and our observations of mutant hatchlings indicate that the ZRS-sZRS deletion does not affect significantly the development of the anal fin. For each stage represented in a-f, five or more individuals were analyzed. Scale bars, 100μm.

  6. Supplementary Figure 6 Detailed analyses of the dorsal fin phenotype in the ZRS ∆948 mutant.

    Bone and cartilage staining showing the dorsal fin from wild type (a) or ∆948 (b-d) 4 month old fish. 74% (31/42) of the mutants show complete ablation (black arrow) of the dorsal fin (d) while in 19% (8/42) no fin rays are observed and endoskeletal elements are highly reduced (c). Only 7% (3/42) of the mutants analysed show fin rays although severely affected (b). Bone and cartilage staining protocol was performed in three independent experiments. Scale bars, 1mm.

  7. Supplementary Figure 7 ZRS and sZRS enhancer expression and skeletal architecture for anal, dorsal and pelvic fins in wild type and ∆401 and ∆948 ZRS mutants.

    a-c, whole mount ISH showing ZRS expression in the posterior region of the anal, dorsal (black arrows) and pelvic fin (white arrows) buds in zebrafish. d-f, whole mount ISH showing sZRS expression in pelvic fins (white arrows), but not in anal or dorsal zebrafish fin buds. ZRS and sZRS expression assessment in a-f was performed in three independent experiments with similar results. Scale bars, 100μm. g-o, Comparative analysis of skeletal elements, as revealed by alcian blue/alizarin red staining, for each of the fins in wild type and mutant adult fish. Ectopic elements are observed in pelvic mutant fins (black arrowheads). Bone staining was performed in three or more independent experiments for wildtype, ∆401 and ∆948 ZRS mutant animals with similar results. Scale bars, 1mm. p-q, Quantitative analysis of pelvic fins length illustrate the phenotypic defects observed in these appendages in the ∆948 mutants (WT mean= 0.088, ∆948 mean=0.041), p-value=1.433x10−5 (***) . In contrast, no significant defects in structure and length are observed for the anal fin of ∆948 mutants (WT mean=0.239, ∆948 mean=0.248, p-value=0.512). Each point in p and q graphs represents measurements from single adult fins. Differences in fin length in p and q were analysed using a t-test.

Supplementary information

  1. Supplementary Figures

    Supplementary Figures 1–7 and Supplementary Table 1

  2. Life Sciences Reporting Summary

  3. Supplementary Table 1

    Oligonucleotides used for PCR amplification of the ZRS and sZRS enhancer from different species. Also listed are primers used for the 4C-seq analyses, genomic deletion target sites (PAM sequence in bold), genomic deletions screening and lamprey cloning.

  4. Supplementary Video 1

    ZRS-sZRS double mutants lack pectoral fins. Video showing a ∆3.4-kb larva (orange arrow) and its siblings freely swimming in a Petri dish recently after hatching (9 dpf).