Migratory neuronal progenitors arise from the neural plate borders in tunicates

Abstract

The neural crest is an evolutionary novelty that fostered the emergence of vertebrate anatomical innovations such as the cranium and jaws1. During embryonic development, multipotent neural crest cells are specified at the lateral borders of the neural plate before delaminating, migrating and differentiating into various cell types. In invertebrate chordates (cephalochordates and tunicates), neural plate border cells express conserved factors such as Msx, Snail and Pax3/7 and generate melanin-containing pigment cells2,3,4, a derivative of the neural crest in vertebrates. However, invertebrate neural plate border cells have not been shown to generate homologues of other neural crest derivatives. Thus, proposed models of neural crest evolution postulate vertebrate-specific elaborations on an ancestral neural plate border program, through acquisition of migratory capabilities and the potential to generate several cell types5,6,7. Here we show that a particular neuronal cell type in the tadpole larva of the tunicate Ciona intestinalis, the bipolar tail neuron, shares a set of features with neural-crest-derived spinal ganglia neurons in vertebrates. Bipolar tail neuron precursors derive from caudal neural plate border cells, delaminate and migrate along the paraxial mesoderm on either side of the neural tube, eventually differentiating into afferent neurons that form synaptic contacts with both epidermal sensory cells and motor neurons. We propose that the neural plate borders of the chordate ancestor already produced migratory peripheral neurons and pigment cells, and that the neural crest evolved through the acquisition of a multipotent progenitor regulatory state upstream of multiple, pre-existing neural plate border cell differentiation programs.

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Figure 1: Bipolar tail neurons come from the borders of the neural plate.
Figure 2: Bipolar tail neuron precursors delaminate and migrate.
Figure 3: Bipolar tail neuron specification and differentiation.
Figure 4: Synaptic connections of bipolar tail neurons.

Change history

  • 18 November 2015

    Minor changes were made to Fig. 4 (arrows), ED Fig. 2b and the ED Fig. 2 legend.

References

  1. 1

    Bronner, M. E. & LeDouarin, N. M. Evolution and development of the neural crest: an overview. Dev. Biol. 366, 2–9 (2012)

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  2. 2

    Yu, J.-K., Meulemans, D., McKeown, S. J. & Bronner-Fraser, M. Insights from the amphioxus genome on the origin of vertebrate neural crest. Genome Res. 18, 1127–1132 (2008)

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  3. 3

    Wada, H., Holland, P. W. H., Sato, S., Yamamoto, H. & Satoh, N. Neural tube is partially dorsalized by overexpression of HrPax-37: the ascidian homologue of Pax-3 and Pax-7. Dev. Biol. 187, 240–252 (1997)

    CAS  Article  Google Scholar 

  4. 4

    Abitua, P. B., Wagner, E., Navarrete, I. A. & Levine, M. Identification of a rudimentary neural crest in a non-vertebrate chordate. Nature 492, 104–107 (2012)

    CAS  ADS  PubMed  PubMed Central  Article  Google Scholar 

  5. 5

    Wada, H. Origin and evolution of the neural crest: a hypothetical reconstruction of its evolutionary history. Dev. Growth Differ. 43, 509–520 (2001)

    CAS  Article  Google Scholar 

  6. 6

    Baker, C. V. H. & Bronner-Fraser, M. The origins of the neural crest. Part II: an evolutionary perspective. Mech. Dev. 69, 13–29 (1997)

    CAS  Article  Google Scholar 

  7. 7

    Shimeld, S. M. & Holland, P. W. H. Vertebrate innovations. Proc. Natl Acad. Sci. USA 97, 4449–4452 (2000)

    CAS  ADS  Article  Google Scholar 

  8. 8

    Jeffery, W. R., Strickler, A. G. & Yamamoto, Y. Migratory neural crest-like cells form body pigmentation in a urochordate embryo. Nature 431, 696–699 (2004)

    CAS  ADS  Article  Google Scholar 

  9. 9

    Mazet, F. et al. Molecular evidence from Ciona intestinalis for the evolutionary origin of vertebrate sensory placodes. Dev. Biol. 282, 494–508 (2005)

    CAS  Article  Google Scholar 

  10. 10

    Pasini, A. et al. Formation of the ascidian epidermal sensory neurons: insights into the origin of the chordate peripheral nervous system. PLoS Biol. 4, e225 (2006)

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  11. 11

    Kaltenbach, S. L., Yu, J.-K. & Holland, N. D. The origin and migration of the earliest-developing sensory neurons in the peripheral nervous system of amphioxus. Evol. Dev. 11, 142–151 (2009)

    Article  Google Scholar 

  12. 12

    Imai, J. H. & Meinertzhagen, I. A. Neurons of the ascidian larval nervous system in Ciona intestinalis: II. Peripheral nervous system. J. Comp. Neurol. 501, 335–352 (2007)

    CAS  Article  Google Scholar 

  13. 13

    Ma, Q., Fode, C., Guillemot, F. & Anderson, D. J. NEUROGENIN1 and NEUROGENIN2 control two distinct waves of neurogenesis in developing dorsal root ganglia. Genes Dev. 13, 1717–1728 (1999)

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  14. 14

    Coric, T., Passamaneck, Y. J., Zhang, P., Di Gregorio, A. & Canessa, C. M. Simple chordates exhibit a proton-independent function of acid-sensing ion channels. FASEB J. 22, 1914–1923 (2008)

    CAS  Article  Google Scholar 

  15. 15

    Aniello, F. et al. Identification and developmental expression of Ci-msxb: a novel homologue of Drosophila msh gene in Ciona intestinalis. Mech. Dev. 88, 123–126 (1999)

    CAS  Article  Google Scholar 

  16. 16

    Wada, S. & Saiga, H. Cloning and embryonic expression of Hrsna, a snail family gene of the ascidian Halocynthia roretzi: implication in the origins of mechanisms for mesoderm specification and body axis formation in chordates. Dev. Growth Differ. 41, 9–18 (1999)

    CAS  Article  Google Scholar 

  17. 17

    Hashimoto, H., Robin, F. B., Sherrard, K. M. & Munro, E. M. Sequential contraction and exchange of apical junctions drives zippering and neural tube closure in a simple chordate. Dev. Cell 32, 241–255 (2015)

    CAS  Article  Google Scholar 

  18. 18

    Nicol, D. & Meinertzhagen, I. Development of the central nervous system of the larva of the ascidian, Ciona intestinalis L: II. Neural plate morphogenesis and cell lineages during neurulation. Dev. Biol. 130, 737–766 (1988)

    CAS  Article  Google Scholar 

  19. 19

    Nakamura, M. J., Terai, J., Okubo, R., Hotta, K. & Oka, K. Three-dimensional anatomy of the Ciona intestinalis tailbud embryo at single-cell resolution. Dev. Biol. 372, 274–284 (2012)

    CAS  Article  Google Scholar 

  20. 20

    Tang, W. J., Chen, J. S. & Zeller, R. W. Transcriptional regulation of the peripheral nervous system in Ciona intestinalis. Dev. Biol. 378, 183–193 (2013)

    Article  CAS  Google Scholar 

  21. 21

    Theveneau, E. & Mayor, R. Neural crest delamination and migration: from epithelium-to-mesenchyme transition to collective cell migration. Dev. Biol. 366, 34–54 (2012)

    CAS  Article  Google Scholar 

  22. 22

    Torrence, S. & Cloney, R. Nervous system of ascidian larvae: caudal primary sensory neurons. Zoomorphology 99, 103–115 (1982)

    Article  Google Scholar 

  23. 23

    Maksimovic, S. et al. Epidermal Merkel cells are mechanosensory cells that tune mammalian touch receptors. Nature 509, 617–621 (2014)

    CAS  ADS  PubMed  PubMed Central  Article  Google Scholar 

  24. 24

    Morrison, K. M., Miesegaes, G. R., Lumpkin, E. A. & Maricich, S. M. Mammalian Merkel cells are descended from the epidermal lineage. Dev. Biol. 336, 76–83 (2009)

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  25. 25

    Horie, T., Kusakabe, T. & Tsuda, M. Glutamatergic networks in the Ciona intestinalis larva. J. Comp. Neurol. 508, 249–263 (2008)

    CAS  Article  Google Scholar 

  26. 26

    Artinger, K. B., Chitnis, A. B., Mercola, M. & Driever, W. Zebrafish narrowminded suggests a genetic link between formation of neural crest and primary sensory neurons. Development 126, 3969–3979 (1999)

    CAS  Google Scholar 

  27. 27

    Fritzsch, B. & Northcutt, R. G. Cranial and spinal nerve organization in amphioxus and lampreys: evidence for an ancestral craniate pattern. Acta Anat. (Basel) 148, 96–109 (1993)

    CAS  Article  Google Scholar 

  28. 28

    Paukert, M. et al. A family of acid-sensing ion channels from the zebrafish: widespread expression in the central nervous system suggests a conserved role in neuronal communication. J. Biol. Chem. 279, 18783–18791 (2004)

    CAS  Article  Google Scholar 

  29. 29

    O’Brien, G. S. et al. Coordinate development of skin cells and cutaneous sensory axons in zebrafish. J. Comp. Neurol. 520, 816–831 (2012)

    PubMed  PubMed Central  Article  Google Scholar 

  30. 30

    Buitrago-Delgado, E., Nordin, K., Rao, A., Geary, L. & LaBonne, C. Shared regulatory programs suggest retention of blastula-stage potential in neural crest cells. Science 348, 1332–1335 (2015)

    CAS  ADS  PubMed  PubMed Central  Article  Google Scholar 

  31. 31

    Stolfi, A. et al. Early chordate origins of the vertebrate second heart field. Science 329, 565–568 (2010)

    CAS  ADS  PubMed  PubMed Central  Article  Google Scholar 

  32. 32

    Russo, M. T. et al. Regulatory elements controlling Ci-msxb tissue-specific expression during Ciona intestinalis embryonic development. Dev. Biol. 267, 517–528 (2004)

    CAS  Article  Google Scholar 

  33. 33

    Stolfi, A. & Christiaen, L. Genetic and genomic toolbox of the chordate Ciona intestinalis. Genetics 192, 55–66 (2012)

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  34. 34

    Khoueiry, P. et al. A cis-regulatory signature in ascidians and flies, independent of transcription factor binding sites. Curr. Biol. 20, 792–802 (2010)

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  35. 35

    Takamura, K., Minamida, N. & Okabe, S. Neural map of the larval central nervous system in the ascidian Ciona intestinalis. Zoolog. Sci. 27, 191–203 (2010)

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  36. 36

    Imai, K. S., Stolfi, A., Levine, M. & Satou, Y. Gene regulatory networks underlying the compartmentalization of the Ciona central nervous system. Development 136, 285–293 (2009)

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  37. 37

    Rothbächer, U., Bertrand, V., Lamy, C. & Lemaire, P. A combinatorial code of maternal GATA, Ets and β-catenin-TCF transcription factors specifies and patterns the early ascidian ectoderm. Development 134, 4023–4032 (2007)

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  38. 38

    Dynes, J. L. & Ngai, J. Pathfinding of olfactory neuron axons to stereotyped glomerular targets revealed by dynamic imaging in living zebrafish embryos. Neuron 20, 1081–1091 (1998)

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  39. 39

    Satou, Y. et al. A cDNA resource from the basal chordate Ciona intestinalis. Genesis 33, 153–154 (2002)

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  40. 40

    Roure, A. et al. A multicassette Gateway vector set for high throughput and comparative analyses in Ciona and vertebrate embryos. PLoS ONE 2, e916 (2007)

    ADS  PubMed  PubMed Central  Article  CAS  Google Scholar 

  41. 41

    Stolfi, A., Wagner, E., Taliaferro, J. M., Chou, S. & Levine, M. Neural tube patterning by Ephrin, FGF and Notch signaling relays. Development 138, 5429–5439 (2011)

    CAS  Article  Google Scholar 

  42. 42

    Davidson, B., Shi, W., Beh, J., Christiaen, L. & Levine, M. FGF signaling delineates the cardiac progenitor field in the simple chordate, Ciona intestinalis. Genes Dev. 20, 2728–2738 (2006)

    CAS  Article  Google Scholar 

  43. 43

    Hudson, C. & Yasuo, H. A signalling relay involving Nodal and Delta ligands acts during secondary notochord induction in Ciona embryos. Development 133, 2855–2864 (2006)

    CAS  Article  Google Scholar 

  44. 44

    Christiaen, L., Wagner, E., Shi, W. & Levine, M. The sea squirt Ciona intestinalis. Cold Spring Harb. Protoc. pdb.emo138 (2009)

  45. 45

    Beh, J., Shi, W., Levine, M., Davidson, B. & Christiaen, L. FoxF is essential for FGF-induced migration of heart progenitor cells in the ascidian Ciona intestinalis. Development 134, 3297–3305 (2007)

    CAS  Article  Google Scholar 

  46. 46

    Ikuta, T. & Saiga, H. Dynamic change in the expression of developmental genes in the ascidian central nervous system: revisit to the tripartite model and the origin of the midbrain–hindbrain boundary region. Dev. Biol. 312, 631–643 (2007)

    CAS  Article  Google Scholar 

  47. 47

    Ando, R., Hama, H., Yamamoto-Hino, M., Mizuno, H. & Miyawaki, A. An optical marker based on the UV-induced green-to-red photoconversion of a fluorescent protein. Proc. Natl Acad. Sci. USA 99, 12651–12656 (2002)

    CAS  ADS  Article  Google Scholar 

  48. 48

    Razy-Krajka, F. et al. Collier/OLF/EBF-dependent transcriptional dynamics control pharyngeal muscle specification from primed cardiopharyngeal progenitors. Dev. Cell 29, 263–276 (2014)

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  49. 49

    Nishida, H. Cell division pattern during gastrulation of the ascidian, Halocynthia roretzi. Dev. Growth Differ. 28, 191–201 (1986)

    Article  Google Scholar 

  50. 50

    Bone, Q. The central nervous system in amphioxus. J. Comp. Neurol. 115, 27–64 (1960)

    Article  Google Scholar 

Download references

Acknowledgements

The authors would like to thank F. Razy-Krajka for assistance with Kaede photoconversion and comments on the manuscript, T. Tolkin for constructing the Mrf reporter plasmid, Z. Lu for ultramicrotomy, and C. Desplan, A. Di Gregorio and all members of the Christiaen and Meinertzhagen labs for feedback and suggestions. We thank H. Hashimoto, F. Robin and N. Takatori for embryo illustration template files. This work was funded by a National Science Foundation Postdoctoral Fellowship in Biology (under grant NSF-1161835) to A.S., by National Institutes of Health award GM096032 to L.C., and by grant DIS0000065 from NSERC (Ottawa) to I.A.M.

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Authors

Contributions

A.S., K.R., I.A.M. and L.C. designed the study, analysed the data, and wrote the paper. A.S. and K.R. performed the experiments.

Corresponding author

Correspondence to Lionel Christiaen.

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The authors declare no competing financial interests.

Extended data figures and tables

Extended Data Figure 1 In situ hybridization of neural plate border markers Snail and Msx.

a, Immunolabelling for β-galactosidase (red) and in situ hybridization for Snail mRNA (green) in stage 12 embryo electroporated with Msx>lacZ, revealing Snail expression in the BTN progenitors (b9.36 cells, arrowheads). Dashed area enlarged in a′. b, Double in situ hybridization for Snail (green on merged image) and Msx (red on merged image) in stage 12 embryos counterstained with DAPI (blue on merged image), showing co-expression in neural plate border cells, including BTN progenitors. Scale bars, 25 μm.

Extended Data Figure 2 Lineage tracing of b9.36 descendants.

a, Photoconversion of Kaede::nls driven by the Msx driver was used to follow the cell divisions of the BTN progenitors from the late gastrula stage to the early tailbud stage. Both b10.71 and b10.72 divide once. b11.141 will give rise to a definitive anterior BTN (see Extended Data Fig. 4). Numbers in each panel represent time in minutes elapsed from the initial photoconversion event. Scale bar, 50 μm. b, Lineage tree showing specification of aBTNs in relation to other cells of the posterior neural plate borders. For simplicity, only one side of the embryo is depicted. c, Lateral view of a 110-cell-stage embryo showing the positions of blastomeres in b. Red lines connect sibling cells. d, Dorsal view of a neurula-stage embryo showing zippering of posterior neural-plate-border-derived capstone cells18 as neural tube closure is initiated. Panels b and d are courtesy of H. Hashimoto and F. Robin (University of Chicago) and N. Takatori (Tokyo Metropolitan University), and partially modelled after ref. 17. Panel c modelled after ref. 49.

Extended Data Figure 3 Neurog cis-regulatory sequences.

a, Schematic diagram representing Neurog locus and 5′ cis-regulatory sequences including b-line and b-line minimal cis-regulatory modules. Peaks represent nucleotide sequence conservation with Ciona savignyi genome. b, Late gastrula embryo (stage 13) electroporated with full-length Neurog (blue) and Nodal b-line (red) reporter constructs. Reporter co-expression is seen in b9.36 descendants on either side of the neural plate. Neurog expression also marks tail-tip lineages of uncertain provenance, previously reported to be descended from b8.21 (ref. 10). Scale bar, 25 μm. c, Neurog b-line reporter. d, Neurog b-line minimal reporter. Scale bars in c, d, 50 μm.

Extended Data Figure 4 Spatiotemporal restriction of Neurog expression.

a, Lateral view of in situ hybridization (ISH) for Neurog (green) in embryo electroporated with Neurog b-line>H2B::mCherry (red) shows that Neurog expression is selectively maintained in only a subset of initially Neurog-expressing neural plate border cells. a′, In the b9.36 lineage, the anterior-most cell (b11.141, solid arrowhead) is always the sole one to express Neurog at this stage, and will go on to become the anterior BTN. Dashed arrowhead indicates b11.142, the sister cell of b11.141, which has downregulated Neurog relative to its sibling. b, b′, The identities of the cells in the tail tip (presumed b8.21-derived) lineages are unclear, but Neurog is similarly restricted (arrowheads) to a single cell on either side of the midline, which we interpret as the definitive posterior BTNs. c, Control embryo treated with DMSO vehicle, showing wild-type pattern of Neurog expression only in b11.141. d, Neurog is expanded to b11.142 upon treatment with the MEK inhibitor U0126 at 7 h.p.f. This condition also results in specification of supernumerary BTNs, presumably due to expanded Neurog expression (see text for details). Thus, downregulation of Neurog in b11.142 also requires MEK/ERK signalling. e, Diagram of the aBTN lineage, descended from the b8.18 blastomere. Scale bars in a, b, 25 μm. Scale bars in c, d, 10 μm.

Extended Data Figure 5 Perturbation of Notch signalling does not alter Neurogenin expression or bipolar tail neuron specification and differentiation.

a, Top, lateral view of a stage 23 embryo electroporated with Msx>H2B::mCherry (magenta nuclei), Neurog b-line>unc-76::eGFP (green) and Msx>nls::lacZ, serving as the wild-type control condition. Bottom, embryo electroporated with same reporters as upper panel, plus Msx>Su(H)-DBM, which encodes a DNA-binding mutant form of the Notch co-activator Rbpj. No discernable difference in Neurog activation or BTN specification was observed between control and Su(H)-DBM conditions (1 of 32 versus 2 of 42 embryos showing ectopic Neurog+ BTNs, respectively). b, Late overexpression of Su(H)-DBM using the Neurog b-line driver similarly did not alter BTN specification/differentiation, as monitored by Asic>unc-76::eGFP reporter expression (0 of 50 control versus 0 of 50 Su(H)-DBM embryos showed ectopic Asic+ BTNs). Scale bars, 50 μm.

Extended Data Figure 6 Cell polarity and morphogenesis of bipolar tail neurons.

a, Embryo at 11.5 h.p.f. (18 °C) with BTNs displaced from clonally related epidermal cells (epid.) labelled by UNC-76::VenusYFP (red), Galnt7ΔC::CFP (green), and H2B::mCherry (blue) driven by Neurog b-line cis-regulatory module. Targeted localization of CFP by the Galnt7 N-terminal signal sequence reveals polarized subcellular distribution of Golgi apparatus on posterior side of BTN nuclei as migration and proximal process extend in an anterior direction. This is distinct from the apical (dorsal) location of the Golgi apparatus in epidermal cells. b, Embryo at 12.5 h.p.f. (18 °C) showing 180° inversion of Golgi apparatus localization to the anterior side of the nucleus, immediately preceding distal process extension. Scale bars in a, b, 50 μm. c, Still frames from a confocal image stack time lapse movie (Supplementary Video 4) showing inversion of Golgi complex (Galnt7ΔC::VenusYFP, green) relative to nuclei (H2B::mCherry, red) in migrating BTNs. Time lapse imaging initiated at 11.5 h.p.f. (18 °C). Time in minutes elapsed from start shown at bottom right of each panel. Anterior BTN (aBTN) indicated by magenta arrowhead, posterior BTN (pBTN) indicated by white arrowhead. Scale bar, 25 μm. d, Diagram showing correlation of average length of proximal (left) and distal (right) processes and angle of Golgi apparatus location relative to cell nucleus along the anterior–posterior axis in BTNs at different time points. Locations of Golgi apparatus represented by rose plots of bins of 20° spanning anterior (0°) and posterior (180°) endpoints around dorsal edge of BTN nucleus. Bin diameters indicate number of cells. Embryos analysed belong to the same pool as embryos in a and b. See Supplementary Table 1 for source data.

Extended Data Figure 7 Proposed evolution of neural crest through the acquisition of multipotency by neural plate border cells.

a, Cartoon diagram depicting a hypothetical path for neural plate border and neural crest evolution, starting with the reconstructed last common olfactorean ancestor, which could have had neural plate borders lined with committed progenitor cells giving rise to several pigmented ocelli and BTN-like peripheral neurons, a condition that may be conserved in extant cephalochordates50. These cells would have been reduced in the highly miniaturized embryos of extant tunicates, while vertebrates are proposed to have co-opted a mesenchymal, multipotency program to bestow these cells with the potential to give rise to pigment cells, peripheral neurons or other derivatives, after a prolonged period of EMT and migration. b, Diagram representing idealized cell lineages in the neural plate borders of tunicate and hypothetical urolfactorean ancestor, in which segregated lineages at the neural plate borders give rise to committed pigment cell or peripheral neuronal progenitors. c, Diagram of simplified neural crest cell lineage deploying a multipotency program downstream of neural plate border specification and upstream of cell differentiation. Thus, neural crest cells could have evolved through redeployment of a multipotency program (intercalation hypothesis)1, or through its maintenance from earlier embryonic stages (heterochrony hypothesis)30.

Extended Data Table 1 Synaptic input from bipolar tail neurons to motor neurons, identified by electron microscopy

Supplementary information

Supplementary Table 1

This table contains Golgi apparatus repositioning and bipolar tail neuron process extension source data. It contains all measurements, when possible, of Golgi apparatus angle relative to nucleus (measure in degrees), and proximal and distal process lengths (measured in μm), for anterior (aBTN) and posterior (pBTN) bipolar tail neuron precursors on a single side (right or left, not indicated) of each embryo imaged. Embryos were grown to three different stages (11.5, 12.5, and 13.5 hours post-fertilization). See main text, methods, and Extended Data Figure 6 for more details. (XLSX 18 kb)

Delaminating, migrating bipolar tail neuron precursors

Time-lapse confocal imaging of embryo electroporated with Neurog b-line>unc-76::VenusYFP (green), imaged starting at 9.5 hours post-fertilization (at 18 °C). Confocal Z-stacks were acquired every 3 minutes for roughly 2 hours and 20 minutes. (MOV 336 kb)

Migrating posterior bipolar tail neuron precursor

Time-lapse confocal imaging of embryo electroporated with and Fog>H. sapiens CD4::eGFP (green), imaged starting at 11 hours, 21 minutes post-fertilization (at 18°C). Confocal Z-stacks were acquired every 2 minutes, for roughly 1 hour, 45 min. The bipolar tail neuron and related epidermis midline cells are labelled by Neurog b-line>unc-76::mCherry (red), while the cell membranes of the entire epidermis is labeled by Fog>H. sapiens CD4::eGFP. (MOV 423 kb)

3D projection of bipolar tail neuron precursor situated on paraxial mesoderm

A 3D confocal projection of embryo imaged in Figure 2c, showing a Neurog reporter-labeled bipolar tail neuron (magenta) migrating along Mrf reporter-labeled paraxial mesoderm-derived muscles (green). The embryo was electroporated only on the right side. Cell outlines are stained by Alexa Fluor 633 phalloidin. (MOV 2126 kb)

Golgi apparatus repositioning in migrating bipolar tail neuron precursors

Time-lapse confocal imaging of embryo electroporated with Neurog b-line>H2B::mCherry (red) and Neurog b-line>Galnt7ΔC::VenusYFP (green), imaged starting at 11.5 hours post-fertilization (at 18°C). Confocal Z-stacks were acquired every minute for roughly 1 hour. (MOV 182 kb)

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Stolfi, A., Ryan, K., Meinertzhagen, I. et al. Migratory neuronal progenitors arise from the neural plate borders in tunicates. Nature 527, 371–374 (2015). https://doi.org/10.1038/nature15758

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