Wnt11 acts on dermomyotome cells to guide epaxial myotome morphogenesis

  1. Ann Kathrin Heilig
  2. Ryohei Nakamura
  3. Atsuko Shimada
  4. Yuka Hashimoto
  5. Yuta Nakamura
  6. Joachim Wittbrodt
  7. Hiroyuki Takeda  Is a corresponding author
  8. Toru Kawanishi  Is a corresponding author
  1. Department of Biological Sciences, Graduate School of Science, University of Tokyo, Japan
  2. Centre for Organismal Studies, Heidelberg University, Germany
  3. Heidelberg Biosciences International Graduate School, Germany

Abstract

The dorsal axial muscles, or epaxial muscles, are a fundamental structure covering the spinal cord and vertebrae, as well as mobilizing the vertebrate trunk. To date, mechanisms underlying the morphogenetic process shaping the epaxial myotome are largely unknown. To address this, we used the medaka zic1/zic4-enhancer mutant Double anal fin (Da), which exhibits ventralized dorsal trunk structures resulting in impaired epaxial myotome morphology and incomplete coverage over the neural tube. In wild type, dorsal dermomyotome (DM) cells reduce their proliferative activity after somitogenesis. Subsequently, a subset of DM cells, which does not differentiate into the myotome population, begins to form unique large protrusions extending dorsally to guide the epaxial myotome dorsally. In Da, by contrast, DM cells maintain the high proliferative activity and mainly form small protrusions. By combining RNA- and ChIP-sequencing analyses, we revealed direct targets of Zic1, which are specifically expressed in dorsal somites and involved in various aspects of development, such as cell migration, extracellular matrix organization, and cell-cell communication. Among these, we identified wnt11 as a crucial factor regulating both cell proliferation and protrusive activity of DM cells. We propose that dorsal extension of the epaxial myotome is guided by a non-myogenic subpopulation of DM cells and that wnt11 empowers the DM cells to drive the coverage of the neural tube by the epaxial myotome.

Editor's evaluation

This study addresses an interesting and underexplored question in developmental biology, specifically cell migration and muscle development. It builds upon prior analysis of the medaka Double anal fin (Da) mutants by using detailed bioinformatic and time-lapse analysis to explain dorsal somite extension. The authors show that dorsal muscle morphogenesis is actively guided by dorsal dermomyotome cells, rather than being passively shaped by physical constraints alone. Looking downstream of Da, they show that Wnt signaling is central to dorsal extension of the epaxial myotome and propose that similar functions may shape the dorsal musculature across vertebrates.

https://doi.org/10.7554/eLife.71845.sa0

Introduction

Active locomotion, which is powered by skeletal muscles in vertebrates, is critical for animals to survive. Vertebrate skeletal muscles consist of axial muscles (head, trunk, and tail muscles) and appendicular muscles (limb muscles). Axial muscles first arose in the chordates to stabilize and enable side-to-side movement of the body axis. In jawed vertebrates, the subdivision of axial muscles into epaxial (dorsal) and hypaxial (ventral) muscles led to an increased range of movement: dorsoventral undulation in fish and lateral movements in terrestrial vertebrates (Goodrich, 2004; Romer and Parsons, 1986; Fetcho, 1987; Sefton and Kardon, 2019). Among these muscles, epaxial muscles are characterized by their unique anatomical structure, which extends dorsally and surrounds the vertebrae. This morphology also ensures mechanical support and protection of the vertebrae and the spinal cord inside. While we have a detailed understanding of how the myotome, precursors of epaxial and hypaxial muscles, differentiates from a somitic compartment called the dermomyotome (DM) (Kalcheim et al., 1999; Gros et al., 2009; Hollway and Currie, 2005), we only begin to understand the cellular and molecular mechanisms of the subsequent morphogenetic processes generating the epaxial muscles. Previous studies in rats and mice suggested that myocytes of the epaxial myotome do not actively migrate dorsally but are guided by external forces (Deries et al., 2010; Deries et al., 2012). However, what exerts such forces to drive extension of epaxial myotome is still unclear.

Fish have been extensively utilized to study myotome development thanks to the transparency of their bodies throughout embryonic development (Wolff et al., 2003; Stellabotte et al., 2007; Nguyen et al., 2017; Ganassi, 2018). Additionally, their epaxial trunk muscles have a simple structure consisting of only one anatomical unit (reviewed in Sefton and Kardon, 2019). Like other vertebrates, fish myotomes, on either side of the neural tube, extend dorsally after somite differentiation and eventually cover the neural tube by the end of embryonic development (Figure 1A). The spontaneous medaka (Oryzias latipes) mutant Double anal fin (Da) displays a particular epaxial myotome morphology, in which the dorsal ends of the left and right epaxial myotome fail to extend sufficiently and thus do not cover the neural tube at the end of embryonic development. Previous studies demonstrated that the dorsal trunk region of the Da mutant is transformed into the ventral one, including not only the myotome but also the body shape, skeletal elements, pigmentation, and fin morphology (Figure 1B and D; Ishikawa, 1990; Ohtsuka et al., 2004). Given the unique morphological features, the medaka Da mutant is an excellent model to study the morphogenesis of epaxial myotome. Genetic analysis of the Da mutant revealed that this phenotype is due to a dramatic reduction of the expression of the transcription factors zic1 and zic4 in the dorsal somites (Figure 1C and E), and identified zic1/zic4 as master regulators of trunk dorsalization (Ohtsuka et al., 2004; Kawanishi et al., 2013). The down-regulation of zic1/zic4 specifically in the dorsal somites is caused by the insertion of a large transposon, disrupting the dorsal somite enhancer of zic1/zic4 (Moriyama et al., 2012; Inoue et al., 2017). While the function and downstream targets of Zic1 and Zic4 have been studied in nervous system development and somitogenesis (Li, 2006; Pan et al., 2011; Himeda et al., 2013; Hong and Saint-Jeannet, 2017; Aruga and Millen, 2018), the molecular mechanism of how these Zic genes control dorsal trunk morphogenesis has not been investigated so far.

Figure 1 with 1 supplement see all
The epaxial myotome of the Da mutant fails to cover the neural tube at the end of embryonic development.

(A) Schematic representation of dorsal somite extension which results in the full coverage of the neural tube by the epaxial myotomes at the end of embryonic development. (B) Lateral view of adult Wt medaka. Dorsal, caudal, and anal fins are outlined. (C) Lateral view of whole-mount in situ hybridization against zic1 in a 12 ss (1.7 dpf, stage 23) Wt embryo. zic1 expression can be observed in the brain, neural tissues and the dorsal somites. (D) Lateral view of adult Da mutant. Dorsal, anal and caudal fins are outlined. The dorsal trunk region resembles the ventral trunk region. (E) Lateral view of whole-mount in situ hybridization against zic1 of a 12 ss Da embryo. zic1 expression can be observed in the brain and the neural tissues, but is drastically decreased in the dorsal somites (arrowhead). (F, H) Cross-sections of tail regions of hematoxylin stained 9 dpf embryos. Dorsal ends of myotomes are outlined. In Wt, the left and the right myotome come in close contact at the top of the neural tube and form a gapless muscle layer (F). In the Da mutant, the left and right myotome fail to come in contact at the top of the neural tube (H). (G, I) Dorsal view of whole-mount Phalloidin (magenta) immunostaining labeling the myotome of Wt (G) and Da (I) embryos. Epaxial myotome is outlined, and asterisks label melanophores. The contour of the myotomes was drawn based on the Z-stack images of the dorsal myotomes to avoid ambiguity caused by melanophores. Anterior to the left. (J) Schematic representation of measurements to analyze the distance between the left and the right dorsal tip of the myotome (yellow) and the cross-sectional area of the dorsal myotome. For each measurement, three consecutive optical cross sections of the 10th somite were analyzed and averaged. (K) Distance between the left and right tip of the dorsal myotome 5.5 dpf – 9 dpf (n = 6 and 5 for Wt and Da embryos, respectively at 5.5 dpf (stage 35) (p = 0.097); n = 5 and 5 at 6 dpf (stage 36) (p = 0.075); n = 8 and 8 at 7 dpf (stage 37) (p = 0.0047); n = 5 and 6 at 8 dpf (stage 38) (p = 0.019); n = 7 and 6 at 9 dpf (stage 39) (p = 0.034); median, first and third quartiles are shown). (L) Cross-sectional area of the dorsal somites at 22 ss (2.25 dpf, stage 26; n = 10 somites of 5 Wt embryos, n = 12 somites of 6 Da embryos) (p = 0.038), 7 dpf (n = 10 somites of 5 Wt embryos, n = 10 somites of 5 Da embryos) (p = 0.044) and 9 dpf (n = 8 somites of 4 Wt embryos, n = 6 somites of 3 Da embryos) (p = 0.0019). Median, first and third quartiles are shown. HM, horizontal myoseptum; NT, neural tube. Scale bar = 50 μm. ** p < 0.01, * p < 0.05, ns, not significant.

Here, we describe the morphogenetic process of the formation of the epaxial myotome of the back, which we termed ‘dorsal somite extension’ (Figure 1A). By in vivo time-lapse imaging we uncovered its cellular dynamics; during dorsal somite extension, dorsal DM cells reduce their proliferative activity and subsequently form unique large protrusions extending dorsally, guiding the epaxial myotome towards the top of the neural tube. We also found that these DM cells form a subpopulation that gives rise to non-myotomal cell lineages during embryonic development. In the Da mutant, by contrast, DM cells keep their high proliferative activity and mainly form small protrusions. Mechanistically, we identified a Zic1 downstream-target gene, wnt11 (former wnt11r Postlethwait et al., 2019), as a crucial factor for dorsal somite extension, and demonstrated that Wnt11 regulates cellular behavior of dorsal DM cells by promoting protrusion formation and negatively regulating proliferation. We thus propose an unprecedented process of epaxial myotome morphogenesis driven by a non-myogenic population of DM cells during embryogenesis.

Results

Our previous study showed that zic1 and zic4 expression starts at embryonic stages and persists throughout life (Kawanishi et al., 2013). Phenotypic analysis of homozygous adult Da mutants implies long-term participation of Zic-downstream genes in the formation of dorsal musculatures, which eventually affects the external appearance of the fish adult trunk. Here, we examined the initial phase of this long-term dorsalization process. In the following of the study, we will focus on zic1, since zic1 and zic4 are expressed in an identical fashion with overlapping functions in trunk dorsalization of medaka, and zic4 is expressed slightly weaker than zic1 (Moriyama et al., 2012; Kawanishi et al., 2013).

The dorsal myotome of the Da mutant fails to cover the neural tube

In wild-type (Wt) medaka, the dorsal ends of the myotomes first came in contact at 7 days post fertilization (dpf, stage 37) and formed the tight, thick myotome layer covering the neural tube at the end of embryonic development (9 dpf, stage 39) (Figure 1F–G, Figure 1—figure supplement 1A-F). In the ventralized Da mutant, however, the dorsal ends of the myotomes did not extend sufficiently and failed to cover the neural tube at the end of embryonic development (Figure 1H–I, Figure 1—figure supplement 1G-L). The ends of the ventralized dorsal myotome in the mutant displayed a round shape (not pointed as found in Wt) which resembled the morphology of the ventral myotome.

We wondered if there are other morphological differences between Wt dorsal myotome and the ventralized dorsal myotome of the Da mutant. Indeed, the cross-sectional area in the Da mutant was significantly larger compared to Wt (Figure 1L). Possible explanations for a larger cross-sectional area in the Da mutant could be a larger myofiber diameter or a higher number of myofibers, which make up the myotome. We measured the diameter of dorsal myotome muscle fibers in Wt and Da mutant embryos but could not observe a difference, suggesting that dorsal myotome of the Da mutant has a higher number of myotomal cells (Figure 1—figure supplement 1M).

Proliferative activity of the dorsal DM cells is enhanced in the Da mutant

In fish, as in other vertebrates, the DM gives rise to muscle precursor cells, which ultimately differentiate into myofibers. In medaka the DM is a one cell-thick, Pax3/7-positive cell layer encompassing the myotome (Figure 2A–B’’; Hollway et al., 2007; Abe et al., 2019). A high proliferative activity of the dorsal DM could explain a larger cross-sectional area of the dorsal myotome of the Da mutant. To test this, we performed immunohistochemistry against the mitotic marker phospho-histone H3 (pH3) and the DM marker Pax3/7 on Wt and Da embryos (Figure 2C–E). In both Wt and Da embryos, pH3-positive cells were randomly distributed in the dorsal DM without obvious bias (Figure 2C–D). At the 12-somite stage (12 ss, 1.7 dpf stage 23), when zic1 expression in the somites becomes restricted to the dorsal region (Kawanishi et al., 2013), the number of pH3-positive DM cells per dorsal somite was not significantly different between Wt and Da. Remarkably, from 16 ss (1.8 dpf, stage 24) onwards, the number of pH3-positive cells became reduced in the Wt, whereas in Da, no such reduction was observed (Figure 2E). At 35 ss (3.4 dpf, stage 30), pH3-positive cells increased both in the Wt and Da, but the mutant DM cells were more proliferative (Figure 2E). Immunohistochemistry against another proliferation marker PCNA confirmed these findings (Figure 2—figure supplement 1A-C). The number of pH3-positive cells in the ventral DM was not significantly different in Wt and Da embryos at 22 ss (2.75 dpf, stage 26) (Figure 2—figure supplement 2A). These results suggest that zic1 reduces proliferative activity of the DM, which becomes evident following the confinement of its expression to the dorsal somite region.

Figure 2 with 2 supplements see all
Wt dorsal DM cells show lower proliferative activity after the confinement of zic1 expression to the dorsal somite.

(A) Lateral view of 35 ss (3.4 dpf, stage 30) embryo, 10th somite is positioned in the center. Pax3/7 (green) labels DM cells and Phalloidin (magenta) labels myotome. The horizontal myoseptum (HM) separates the myotome into epaxial myotome (dorsal) and hypaxial myotome (ventral). (B–B’’) Optical cross sections, myotome is labeled by Phalloidin (magenta) and encompassed by a one-cell thick layer of DM labeled with Pax3/7 (green). Asterisks mark neural crest cells which are highly Pax3/7-positive (B’’). (C, D) Lateral view of Wt (C) and Da (D) 35 ss embryos labeled with Pax3/7 (green) and pH3 (magenta; representatives are indicated by arrowheads). Asterisks mark neural crest cells. (E) Quantification of pH3-positive cells in Wt and Da at 12 ss (n = 46.5 somites from 4 Wt embryos, n = 95 somites from 7 Da embryos; p = 0.48), 16 ss (n = 54.5 somites from 5 Wt embryos, n = 42.5 somites from 4 Da embryos; p = 0.0038), 22 ss (n = 66 somites from 6 Wt embryos, n = 40.5 somites from 5 Da embryos; p = 0035) and 35 ss (n = 49 somites from 6 Wt embryos, n = 47.5 somites from 5 Da embryos; p = 0.0008). Median, first and third quartiles are shown. ns, not significant, **p < 0.01, ***p < 0.001. Anterior to the left. Dorsal to the top. DM, dermomyotome; HM, horizontal myoseptum; M, myotome; NT, neural tube; NC, notochord. Scale bar = 50 μm.

Wt dorsal DM cells form numerous large, motile protrusions at the onset of dorsal somite extension

The epaxial myotome, on either side of the neural tube, extends dorsally to cover the neural tube by the end of embryonic development. In the Da mutant, the myotome is unable to cover the neural tube despite increased dorsal myotome growth at the hatching stage (Figures 1L and 2). This suggest that additionally to physical extension an active process might support the dorsal movement of somites. To examine the behavior of zic1-positive cells underlying this dorsal somite extension, we performed in vivo time-lapse imaging of dorsal somites using the transgenic line Tg(zic1:GFP,zic4:DsRed), which expresses GFP under the control of the zic1 promoter and enhancers to visualize the dorsal somitic cells (Kawanishi et al., 2013) (hereafter called Tg(zic1:GFP) since the DsRed fluorescence was negligible in the following analyses). Intriguingly, around 22 ss and onwards, cells at the tip of the dorsal somites started to form numerous large protrusions extending dorsally towards the top of the neural tube (Figure 3A, Video 1). We defined the beginning of protrusion formation as the onset of dorsal somite extension. Close-up views of the time-lapse images (Figure 3A, Video 1) showed that these protrusions were motile and dynamically formed new branches at their dorsal tips (Figure 3J, Figure 3—figure supplement 1D). Immunohistochemistry revealed that the protrusion-forming cells belong to the DM (Figure 3—figure supplement 1A-A'').

Figure 3 with 2 supplements see all
Wt dorsal DM cells form numerous large, motile protrusions at the onset of somite extension.

(A, B) Dorsal view of time-lapse in vivo imaging of onset of dorsal somite extension at 24 ss (2.4 dpf, stage 27) of Tg(zic1:GFP) (A) and Tg(zic1:GFP);Da (B) embryos. 15th somite is positioned in the center, z-stacks were imaged every 10 min, time is displayed in min. Asterisks indicate migrating melanophore. Scale bar = 50 μm. (C, F) Lateral view of in vivo imaging of Tg(zic1:GFP) (C) and Tg(zic1:GFP);Da (F). Signals in dorsal somites are highlighted in green. Arrowheads indicate small protrusions, arrow indicates large protrusion. Scale bar = 50 μm. (D, G) Dorsal view of in vivo imaging of large protrusion in Wt (D) and Da (G). Embryos were injected with Actin-Chromobody GFP (AC-GFP) mRNA. Signals in somitic cells are highlighted in green. Brackets indicate a lamellipodia-like structure. Arrows indicate filopodia branching out from dorsal tips of lamellipodia-like core. Asterisks indicate sclerotome cells. Scale bar = 25 μm. (E, H) Summary of dorsal DM cell protrusions in Wt (E) and Da (H) embryos. Arrowheads indicate small protrusions, brackets indicate lamellipodia-like core structure of large protrusions, and arrows indicate filopodia bundles branching off from dorsal tips of large protrusions. (I-I’’) Quantification of protrusions from Tg(zic1:GFP) (n = 14 embryos) and Tg(zic1:GFP);Da (n = 11 embryos). Protrusions of the 8th-12th somite were counted (mean ± SD, p = 0.01 for small protrusions, p = 3.3e-08 for large protrusions, p = 2.1e-05 for total protrusions). (J–K) Lateral view of protrusions extracted from time-lapse imaging of (A–B). Arrowheads indicate tip of protrusions, time is displayed in min. (J) Protrusion observed in Tg(zic1:GFP). (K) Protrusion observed in Tg(zic1:GFP);Da. Scale bar = 25 μm. (L, M) Z-planes of large protrusions of 24 ss embryos treated with DMSO (L) or ML141 (M). Arrowheads indicate filopodia. Scale bar = 25 μm. (N) Distance between dorsal somite tip and top of neural tube in 24 ss embryos (n = 12 somites from 6 Wt embryos, n = 8 somites from 4 Da embryos, p = 6.8e-05; n = 10 somites from 5 embryos treated with DMSO, n = 14 somites from 7 embryos treated with ML141, p = 0.021). Median, first and third quartiles are shown. ****p < 0.0001, *p < 0.5. Anterior to the left. DM, dermomyotome; M, myotome; NT, neural tube.

Video 1
Onset of dorsal somite extension in Tg(zic1:GFP).

Dorsal view of time-lapse in vivo imaging of 24 ss Tg(zic1:GFP) embryo. 15th somite is positioned in the center, z-stacks were imaged every 10 min, time is displayed in min. Anterior to the left. Scale bar = 50 μm.

To characterize the protrusions, we classified them according to their length into small ( < 8 μm, Figure 3C arrowheads) and large ( ≥ 8 μm, Figure 3C arrow) protrusions (Figure 3—figure supplement 1B-C). Based on their shape, we reasoned that the small protrusions correspond to lamellipodia (Figure 3C, arrowheads), while the large protrusions appeared more complex. To investigate the nature of large protrusions, we injected Actin-Chromobody GFP mRNA to visualize the actin skeleton. The large protrusions were found to exhibit a complex architecture consisting of a lamellipodia-like core structure (Figure 3D, bracket) with additional multiple bundles of filopodia (protrusions with linear arranged actin filaments) branching out from their dorsal tips (Figure 3D, arrows, summarized in Figure 3E).

Interestingly, time-lapse in vivo imaging of Tg(zic1:GFP);Da showed that in the Da background, protrusions started to form later (Figure 3B, Video 2) and the number of large protrusions and protrusions in total was significantly lower than in Wt (Figure 3I–I’’). In addition, protrusions were transient and mostly failed to form new branches at their dorsal tips (Figure 3K, Figure 3—figure supplement 1E). While no difference in the actin skeleton of small protrusions could be observed, filament bundles branching out from large protrusions of Da DM cells contained fewer and shorter filopodia compared to Wt (Figure 3G, arrowheads, summarized in Figure 3H). These results indicate that the protrusive activity, especially the ability to form large protrusions, is significantly reduced in the Da mutant. The large protrusions of the dorsal DM cells continuously appeared at later stages of dorsal somite extension, too (Figure 3—figure supplement 2).

Video 2
Onset of dorsal somite extension in Tg(zic1:GFP);Da.

Dorsal view of time-lapse in vivo imaging of 24 ss Tg(zic1:GFP);Da embryo. 15th somite is positioned in center, z-stacks were imaged every 10 min, time is displayed in min. Bright cell at the bottom migrating to the right is a melanophore. Anterior to the left. Scale bar = 50 μm.

To investigate the role of large protrusions during the onset of dorsal somite extension, we inhibited filopodia formation using ML141 (Figure 3L and M). ML141 specifically inhibits Cdc42/Rac1 GTPases, which are critical for filopodia formation (Hong et al., 2013; Fantin, 2015). Intriguingly, dorsal somites of embryos treated with ML141 extended significantly less dorsally compared to control embryos treated with DMSO (Figure 3N). Taken together, our data suggest that the unique large protrusions of the dorsal DM are involved in guiding the epaxial myotome dorsally and zic1 might promote this function.

DM cells delaminate and accumulate between opposing somites during the late phase of dorsal somite extension

We continued to trace the behavior of the DM tip cells until the somites reach the top of the neural tube. Indeed, in vivo imaging of Tg(zic1:GFP) revealed that dorsal DM cells continued to form protrusions, and additionally, some of them delaminated to become Zic1-positive mesenchymal cells accumulating in the space between the dorsal ends of the left and the right somites (Figure 4A, star-shaped cells, arrowheads) from 4.5 dpf (stage 33) onwards. This is consistent with previous observations of strongly zic1 expressing mesenchymal cells in Wt at late embryonic stages (Ohtsuka et al., 2004). As dorsal somite extension proceeded, the number of these mesenchymal cells increased, filling the space between the two myotomes (Figure 4A–C, arrowheads, Videos 34, arrowheads indicate representative mesenchymal cells). These mesenchymal cells formed protrusions towards neighboring mesenchymal cells and DM cells at the tip of somites, creating a dense cellular network between the dorsal ends of the somites. Mosaic cell-labelling demonstrated that the mesenchymal cells originated from the DM (Figure 4—figure supplement 1A-F, arrowheads). Interestingly, while mesenchymal cells dynamically formed protrusions, they showed no extensive migratory behavior and were rather stationary (Videos 34, Figure 4—figure supplement 2). This could suggest that these protrusions fulfill a non-migratory function. When the opposing somites came in contact with each other at 8 dpf (stage 38, 1 day before hatching), the mesenchymal cells tended to attach to the nearest DM cells at the tip of the somite, bridging the gap between the left and right DM cells (Figure 4I, Figure 4—figure supplement 1M-M'''').

Figure 4 with 3 supplements see all
DM cells delaminate from the dorsal somite at the end of dorsal somite extension and accumulate between somites.

(A–F) Dorsal (top) and cross-sectional (bottom) views of Tg(zic1:GFP) at 4.5 dpf (A), 5 dpf (B), and 6 dpf (C) and Tg(zic1:GFP);Da at 4.5 dpf (D), 5 dpf (E) and 6 dpf (F). 10th somite is positioned in center. Arrowheads point to exemplary mesenchymal DM cells, and asterisks mark melanophores. (G, H) Dorsal view of mesenchymal DM cells of Tg(zic1:GFP) (G) and Tg(zic1:GFP);Da (H). (I) Z-plane of dorsal view of 10th somite of Tg(zic1:GFP) embryo. Mesenchymal DM (mes. DM) cells are colored green. Arrowheads indicate exemplary protrusions formed between somitic DM cell and mesenchymal DM cell. (J-K’) Cross-sections of 9th somite before ablation (J), 8 hr post ablation (hpa) (J’), and 10th somite before ablation (K) and 8 hpa (K’) of Tg(zic1:GFP) 5.5 dpf embryo. Green arrowhead indicates the ablation site, and asterisk indicates a pigment cell. (L) Quantification of relative distance between the left and the right tips of the 9th and 10th somites after laser ablation (n = 6 Tg(zic1:GFP) embryos). (M-O’) Fate mapping, using KikGR-mediated photoconversion, of dorsal DM cells with protrusions. (M) A single dorsal DM cell at the dorsal tip of a somite (arrowhead) was labeled at 3.5 dpf. Cross-section of the trunk is shown. Asterisk indicates an ectopically labeled epidermis cell. (N-O’) At hatching stage (9 dpf), descendants of the labeled DM cell contribute to blood vessels (arrowheads in N, N’) and fin mesenchyme in the dorsal fin fold (arrowheads in O, O’). (N) and (O), lateral views; (N’) and (O’), cross sections. Asterisks indicate autofluorescent pigment cells. Anterior to the left. M, myotome; NT, neural tube; S, somite. Scale bar = 25 μm.

Video 3
Mesenchymal DM cells during dorsal somite extension at 4.5 dpf.

Dorsal view of time-lapse in vivo imaging of 4.5 dpf Tg(zic1:GFP) embryo. 10th somite positioned in center, z-stacks were imaged every 10 min, time is displayed in min. Arrowhead indicates representative mesenchymal DM cell. Anterior to the left. Scale bar = 50 μm.

Video 4
Mesenchymal DM cells during dorsal somite extension at 5.5 dpf.

Dorsal view of time-lapse in vivo imaging of 5.5 dpf Tg(zic1:GFP) embryo. 10th somite is positioned in center, z-stacks were imaged every 10 min, time is displayed in min. Arrowhead indicates representative mesenchymal DM cell. Anterior to the left. Scale bar = 50 μm.

In Da mutants, mesenchymal DM cells were also detected in the space between the two myotomes, but the timing of their appearance was delayed, that is between 5 dpf (stage 34) and 5.5 dpf (stage 35) (Figure 4D–F) (4.5 dpf in Wt). Additionally, Da mesenchymal cells exhibited a rounder morphology and formed significantly fewer protrusions compared to Wt while the number of mesenchymal cells was not largely affected (Figure 4G and H, Figure 4—figure supplement 1G-L).

To examine the function of mesenchymal DM cells, we ablated these cells between the 10th somites of 5.5 dpf Tg(zic1:GFP) embryos using an UV laser (Figure 4J–K’, Figure 4—figure supplement 3A-B'). Intriguingly, the distance between the left and the right tips of the 10th somites increased after ablating the mesenchymal DM cells while the neighboring 9th somites continued to shorten the gap (Figure 4L). This suggests that mesenchymal DM cells hold the left and the right somite together to promote the dorsal somite extension at this late phase of myotome development. After 16 hr post ablation, dorsal extension of the 10th somites eventually resumed as the mesenchymal DM cells regenerated at the ablation site (Figure 4L).

Collectively, dorsal DM cells and the mesenchymal cells derived from them seem to actively participate in the entire process of dorsal somite extension, from its onset to neural tube coverage at the end.

Finally, we examined the fate of the dorsal DM cells that derive the mesenchymal cells. We employed a photoconversion technique mediated by a photoconvertible protein KikGR to specifically label a single dorsal DM cell exhibiting protrusions at the tip of a 10th and 20th somites during dorsal somite extension (Figure 4M, arrowhead) and tracked them until the hatching stage. We found that the labeled dorsal DM cells eventually differentiated into cells surrounding blood vessels (likely mural cells, n = 8/9 and 4/8 for 10th and 20th somites, respectively; Figure 4N and N’, arrowheads) and mesenchymal cells in the dorsal fin fold (n = 4/8 for 20th somites; Figure 4O and O’, arrowheads); however, we did not observe any labeled axial muscles (n = 0/9 and 0/8 for 10th and 20th somites, respectively). This suggests that the dorsal DM cells at the tip of somites, guiding myotome extension, do not become myotomal cells themselves at least until the end of embryonic development (hatching stage, 9 dpf).

Zic1 regulates the expression of dorsal-specific genes during somite differentiation

We then addressed the molecular machinery controlling the dorsal somite extension investigated above. Since somite extension is impaired in the zic1-enhancer mutant Da (Moriyama et al., 2012; Kawanishi et al., 2013), we reasoned that downstream genes of Zic1 are regulators of this process.

First, we identified genes which are specifically expressed in the dorsal somites. For this, we collected somites of the transgenic line Tg(zic1:GFP) by removing other tissues after pancreatin treatment, FACS sorted them into the dorsal (GFP+) and ventral (GFP-) somitic cells including the DM cells, and performed RNA-seq and ATAC-seq on both cell populations (Figure 5A, Figure 5—figure supplement 1A). The RNA-seq identified 1,418 differentially expressed genes. Among them 694 genes showed higher expression in the dorsal somites (termed hereafter dorsal-high genes), and 724 genes showed higher expression in the ventral somites (termed hereafter dorsal-low genes) (Figure 5C). We confirmed that zic1 and zic4 were found among the dorsal-high genes (Figure 5C, Supplementary file 1).

Figure 5 with 2 supplements see all
Zic1 regulates dorsal-specific expression of genes in the somites.

(A) Schematic representation of preparation of dorsal and ventral somite cells for RNA-seq and ATAC-seq. (B) Schematic representation of generating the transgenic line Tg(zic1:zic1-Myc);Da. Somites expressing zic1-Myc are subjected to ChIPmentation against Myc. (C) Analysis of RNA-seq revealed that 694 genes are expressed specifically in the dorsal somites (among these genes are zic1, zic4 and wnt11) and 724 genes are expressed specifically in ventral somites. Magenta indicates differentially expressed genes (adjusted p-value < 0.01). (D) The ChIPmentation against Zic1 revealed that 324 of dorsal-high genes, 192 dorsal-low genes and 2716 non differential expressed genes in the somites are potential Zic1 target genes.

Next, we identified potential direct Zic1-target genes by investigating Zic1 binding sites. Since there were no suitable antibodies available to perform ChIP-seq against medaka Zic1, we created a transgenic line expressing a Myc tagged zic1 in the Da background under the control of zic1 promoter and enhancers (Tg(zic1:zic1-Myc,zic4:DsRed);Da, called Tg(zic1:zic1-Myc);Da hereafter). This transgene (zic1:zic1-Myc) efficiently rescued the ventralized phenotype (dorsal and anal fin shape, pigmentation pattern, and body shape) of the Da mutant (Figure 5—figure supplement 1B-C), verifying the full functionality of the tagged Zic1 protein. Somites of the transgenic line Tg(zic1:zic1-Myc);Da were dissected and subjected to ChIP-seq using antibodies against Myc (Figure 5B). Since somites contain a low number of cells, we applied ChIPmentation (Schmidl et al., 2015) to identify genome-wide Zic1 binding sites. From two biological replicates, 5247 reliable ChIP peaks were identified, and we confirmed that the enriched DNA motif among these peaks was showing high similarity with previously identified binding motifs of ZIC family proteins (Figure 5—figure supplement 2A-B). Then, we associated each Zic1 peak to the nearest gene within 50 kb and identified 3232 genes as Zic1 target genes. By comparing the ChIPmentation with RNA-seq data, we found that Zic1 target genes are overrepresented in dorsal-high and dorsal-low genes. While 47% of the dorsal-high genes and 27% of the dorsal-low genes were found to be Zic1 target genes, only 12% of genes which showed no differential expression in dorsal or ventral somites were potential Zic1 downstream target genes (Figure 5D, Supplementary file 1). This suggests that Zic1 can function as transcriptional activator and repressor of versatile genes, but the former role seems to be dominant.

wnt11 is a direct downstream target of Zic1 and down-regulated in the dorsal somites of the Da mutant

To identify potential regulators of dorsal somite extension, we further investigated the differentially expressed Zic1 target genes. Gene Ontology (GO) analysis indicated that both dorsal-high and dorsal-low gene groups, regardless of whether they are Zic1 targets or non-Zic1 targets, were significantly enriched in development related GO terms (Figure 6A, Figure 6—figure supplement 1A, Supplementary files 2 and 3). This indicates that Zic1 regulates a number of developmental genes both directly and indirectly. These results are consistent with the fact that Zic1 regulates various dorsal-specific morphologies of somite-derivatives (Kawanishi et al., 2013).

Figure 6 with 1 supplement see all
wnt11 is a direct downstream target of Zic1 and down-regulated in the dorsal somites of the Da mutant.

(A) GO analysis of dorsal-high (left) and dorsal-low (right) Zic1 target genes. (B) Pathway enrichment analysis indicated that genes associated with Wnt signaling pathway are specifically enriched in dorsal-high Zic1 target genes. (C) Analysis of the wnt11 locus. Peaks form the ChIPmentation against Zic1 (magenta track) overlap with open chromatin regions in the dorsal somites (black track), while this genomic region is less open in ventral somites (grey track). RNA-seq data revealed that wnt11 is highly expressed in dorsal somites (dark green track) and minimally in ventral somites (light green track). (D, E) Dorsal view of tails of whole-mount in situ hybridization against wnt11 performed in Wt (D) and Da (E) 12 ss embryos. The expression of wnt11 is reduced in the Da mutant. (F) RT-PCR performed on pooled tails of Wt and Da 12 ss embryos indicates that zic1 expression is reduced by 8.8-fold and wnt11 expression by 1.4-fold in the Da mutant. Anterior to the left. Scale bar = 100 μm.

In the dorsal-high Zic1 targets, GO terms related to cell migration showed higher enrichment (e.g. ‘chemotaxis’ (p = 5.62E-10), ‘locomotion’ (6.65E-15), ‘ameboidal-type cell migration’ (p = 9.32E-10)) than dorsal-low genes or non-Zic1 target genes (Supplementary file 3). Interestingly, Wnt signaling pathway genes (e.g.: axin2, wnt11, sp5, lrp5, fzd10, prickle1a) and semaphorin-plexin signaling pathway genes (e.g.: sema3a, sema3c, sema3g, plxna1, plxna2, plxnb2, plxnb3, nrp2) were included in these gene groups. Indeed, the GO terms ‘Wnt signaling pathway’ and ‘Semaphorin-plexin signaling pathway’ themselves were also significantly enriched (p = 4.45E-7 and 1.94E-8, respectively) in dorsal-high Zic1 target genes (Supplementary file 2). This suggests that Zic1 directly regulates Wnt pathway, semaphorin and plexin genes, possibly to regulate cell movement in the dorsal somites. We also noticed that the terms ‘extracellular matrix organization’ (p = 7.74E-7; e.g.: adamts20, fbln1), ‘cell communication’ (p = 1.95E-5; e.g.: efna5, epha3) are enriched, suggesting that these genes also affect dorsal somite cell behavior.

Wnt signaling pathway components also ranked high among the dorsal-high Zic1 target genes by pathway enrichment analysis (Figure 6B, Figure 6—figure supplement 1B). Among them, we focused on wnt11 for further analyses due to the following reasons: First, wnt11 was one of the most differentially expressed genes in dorsal somites (Figure 5C, Supplementary file 1). Second, previous studies implicated Wnt11 in protrusion formation and cell migration (Ulrich et al., 2003; De Calisto et al., 2005; Garriock and Krieg, 2007; Matthews et al., 2008). This is particularly interesting since DM cells also exhibit protrusions and migration activity during dorsal somite extension, which are defective in Da mutants.

At the wnt11 locus, peaks of the Zic1-ChIP overlapped with intergenic open chromatin regions downstream of wnt11. These sites were more accessible in dorsal somites than in ventral somites, suggesting that Zic1 regulates wnt11 directly (Figure 6C). Additionally, in situ hybridizations against wnt11 performed on Wt and Da embryos indicated that wnt11 expression is significantly reduced at the dorsal tip of Da dorsal somites including the DM (Figure 6—figure supplement 1C-D). Already when the zic1 expression becomes restricted in the dorsal somites (12 ss) and with proceeding development when zic1 expression gets further restricted to the most dorsal part of the somites and mesenchymal DM cells (Ohtsuka et al., 2004), the expression of wnt11 in the dorsal somites of the Da mutant was reduced compared to Wt (Figure 6D–F, Figure 6—figure supplement 1C-F').

Taken together, we identified wnt11 as promising downstream target gene of Zic1, which could be a novel somite dorsalization factor and play a role in dorsal somite extension.

Wnt11 regulates cell behavior of dorsal DM

Our previous RNA-seq dataset revealed that the expression of wnt11 starts before gastrulation and increases as development proceeds (Figure 7—figure supplement 1A; Nakamura et al., 2021). Injection of single guide RNAs (sgRNAs) against wnt11 to knock out the gene revealed that the resultant genetically mosaic F0 embryos displayed delayed epiboly movement and subsequent morphological defects, including shorter body axis and impaired trunk development (Figure 7—figure supplement 2), implying the difficulty of assessment of the wnt11 function at later stages using the wnt11 mutant. To circumvent the problem, we took two different approaches to knock-down wnt11 during dorsal somite extension. Firstly, we used a wnt11 anti-sense morpholino (wnt11 MO) and determined a concentration which resulted in maximal knock-down effects during dorsal somite extension with minimal gastrulation phenotypes (see Materials and methods). Secondly, we performed temporally controlled knock-down of wnt11 after gastrulation using photo-cleavable Photo-Morpholinos (PhotoMOs) (Tallafuss, 2012; Figure 7A). Strikingly, in wnt11 Photo-Morphants with severe phenotypes the dorsal myotome failed to cover the neural tube (n = 7, Figure 7B–C), a phenotype similar to the Da mutant myotome. Additionally, the myofibers of the Photo-Morphants at 9 dpf were shorter and less organized. This phenotype is consistent with a previously reported role of Wnt11 during early myogenesis, where it regulates the elongation and orientation of myoblasts (Gros et al., 2009).

Figure 7 with 2 supplements see all
Knock-down of wnt11 in Wt embryos recapitulates the Da dorsal somite phenotype.

(A) Schematic outline of PhotoMO mediated knock-down of wnt11. (B, C) Dorsal view of maximum projection of whole-mount Phalloidin staining of 9 dpf embryos injected with PhotoMO but not photocleaved (B) and photo-cleaved (C). The epaxial myotomes of the wnt11 morphant are affected. (D) Quantification of pH3-positive cells in the dorsal DM of 22 ss embryos. Embryos with reduced levels of wnt11 in their dorsal somites (n = 40.5 somites from 5 Da embryos (adopted from Figure 2E), n = 61.5 somites of 6 wnt11 PhotoMO-Morphant embryos and n = 52 somites from 5 wnt11 morphant embryos) have significantly more pH3-positive cells in their dorsal DM compared to the respective controls (n = 66 somites from 6 Wt embryos (adopted from Figure 2E), n = 51 somites of 5 uninduced PhotoMO embryos and n = 65 somites from 6 Control MO embryos; p = 0.0035, 0.0091, 0.021, respectively). Median, first and third quartiles are shown. (E) Dorsal view of 24 ss Tg(zic1:GFP) embryo injected with wnt11 PhotoMO and photo-cleaved at 4 ss. Z-stacks were recorded every 10 min, time is displayed in min, and 15th somite is positioned in the center. (F, G) Lateral view of dorsal somites (colored magenta) of Tg(zic1:GFP) injected with Control MO (F) and wnt11 MO (G). (H-H’’) Quantification of protrusions formed by the 8th-12th somite of Tg(zic1:GFP) injected with Control MO (n = 10 embryos) or wnt11 MO (n = 13 embryos) (mean ± SD, p = 0.069 for small protrusions, p = 7.7e-08 for large protrusions, p = 0.00064 for total protrusions). Anterior to the left. NT, neural tube. Scale bar = 50 μm. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001, ns, not significant.

Next, we investigated the proliferative activity of dorsal DM cells in wnt11 morphants and found that knock-down of wnt11 (either by conventional MO or by PhotoMO) induced significantly more pH3-positive cells per somite, compared to control embryos at 22 ss (Figure 7D). These findings are similar to the observations previously made in the dorsal DM of Da mutants, and suggest that Wnt11 is negatively regulating proliferation of dorsal DM cells.

We then explored whether the onset of dorsal somite extension in wnt11 morphants is similarly impaired as in the Da by time-lapse in vivo imaging. Remarkably, we observed a protrusion formation behavior of wnt11 morphant DM cells similar to Da DM cells, namely delayed onset of protrusion formation and the formation of fewer, shorter protrusions (Figure 7E, Video 5). Quantification of protrusions indicated that wnt11 morphants have significantly fewer large protrusions and protrusions in total, compared to control embryos (Figure 7F–H’’).

Video 5
Onset of dorsal somite extension in wnt11 morphant embryo.

wnt11 was knocked down using the PhotoMO approach (Figure 7A) in a Tg(zic1:GFP) embryo. Dorsal view of time-lapse in vivo imaging of 24 ss Tg(zic1:GFP) embryo. 15th somite is positioned in the center, z-stacks were imaged every 10 min, time is displayed in min. Anterior to the left. Scale bar = 50 μm.

Overall, knock-down of the Zic1 target gene wnt11 recapitulated the phenotype of Da DM cells (Figure 3G–G’’), suggesting the crucial role of Wnt11 in regulating protrusion formation of DM cells.

To further confirm the importance of Wnt11 during dorsal somite extension, we performed rescue experiments in Da embryos. At 18 ss (2.1 dpf, stage 25), we injected a mix of human recombinant Wnt11 (hrWnt11) protein (or BSA in the control group) and Dextran Rhodamine onto the top of the 10th somite of Tg(zic1:GFP);Da embryos (Figure 8A–B’’). Strikingly, the dorsal DM of Da embryos injected with Wnt11 formed significantly more large protrusions and more protrusions in total compared to Da embryos injected with BSA only (Figure 8C–C’’). This indicates that the ventralized Da DM protrusion phenotype can be partially rescued by Wnt11 protein injections and further emphasizes the importance of Wnt11 in somite dorsalization and during dorsal somite extension.

Wnt11 injections partially rescue the Tg(zic1:GFP);Da protrusion phenotype.

(A) Schematic representation of Wnt11 protein injection on top of the 10th somite of 18 ss Tg(zic1:GFP);Da embryos. (B-B’’) Dorsal view of dorsal somites of Tg(zic1:GFP);Da embryos injected with Wnt11-Dextran Rhodamine mix (magenta). GFP signals in dorsal somites are highlighted in green. Protrusions are outlined. (C-C’’) Quantification of dorsal protrusions of the 9th – 11th somite (somite at injection site plus adjacent somites) of embryos injected with BSA (n = 7 embryos) or hrWnt11(n = 7 embryos) (mean ± SD, p = 0.65 for small protrusions, p = 0.0095 for large protrusions, p = 0.03 for total protrusions). Anterior to the left. NT, neural tube. Scale bar = 50 μm. *p < 0.05, **p < 0.01, ns, not significant.

Wnt11 acts through the Wnt/Ca2+ signaling pathway at the onset of somite extension

Finally, we investigated through which signaling pathway the non-canonical Wnt11 acts. In Xenopus, Wnt11 acts through the Wnt/Ca2+ pathway regulating the migration of cells from the dorsal somite and the neural crest into the dorsal fin fold (Garriock and Krieg, 2007). To examine whether this signaling pathway also plays a role during dorsal somite extension, we inhibited the Wnt/Ca2+ signaling pathway using KN-93 in Tg(zic1:GFP) embryos from 4 ss (1.3 dpf, stage 20)–22 ss (Figure 9A). KN-93 specifically inhibits CaMKII, a component of the Wnt/Ca2+ pathway, by binding to it and preventing its phosphorylation, thus keeping it in an inactivated state (Sumi et al., 1991; Tombes et al., 1995; Wu and Cline, 1998; Garriock and Krieg, 2007; Rothschild et al., 2013). Embryos treated with KN-93 showed a higher number of pH3-positive dorsal DM cells per somite, compared to embryos in the control group (Figure 9B). Furthermore, DM cells at the tip of the dorsal somite of embryos treated with KN-93 formed significantly fewer large and fewer protrusions in total, compared to embryos in the control group (Figure 9C–C’’), although the effect was less significant compared to the wnt11 morphants (Figure 7).

Wnt11 acts through the Wnt/Ca2+ signaling pathway during dorsal somite extension.

(A) Schematic representation of KN-93 treatment, embryos in the control group were treated with DMSO. (B) Embryos treated with KN-93 (n = 65.5 somites from 6 embryos) have significantly more pH3-positive cell in the dorsal DM compared to embryos of the control group (n = 52.5 somites from 5 embryos) (median, first and third quartiles, p = 0.045). (C-C’’) Quantification of protrusions formed by the 8th-12th somites of Tg(zic1:GFP) embryos treated with DMSO (n = 10 embryos) or KN-93 (n = 13 embryos) (mean ± SD, p = 0.65 for small protrusions, p = 0.014 for large protrusions, p = 0.0017 for total protrusions). *p < 0.05, **p < 0.01, ns, not significant.

From these results, we suggested that Wnt11 potentially acts through the Wnt/Ca2+ signaling pathway at the onset of somite extension. Wnt/Ca2+ signaling pathway is known to regulate actin polymerization (Choi and Han, 2002; Kohn and Moon, 2005) which could explain the dynamic protrusive activity of the DM cells (Figure 3), emphasizing the importance of Wnt11 during epaxial myotome morphogenesis.

Discussion

Here, we elucidated the key developmental process underlying the epaxial myotome morphogenesis to cover the neural tube in the teleost, medaka. DM cells have been known to serve as a progenitor pool for myotomal and dermal cells (Ben-Yair and Kalcheim, 2005; Hollway et al., 2007; Stellabotte and Devoto, 2007). In our study, we showed that DM cells at the tip of the dorsal somite form unique large protrusions, guiding the myotome towards the top of the neural tube. Furthermore, these DM cells do not give rise to myotomal cells during embryonic development. We found that zic1 and its direct downstream target wnt11 are crucial factors for this morphogenetic process. We thus revealed a novel role of non-myogenic DM cells during epaxial myotome morphogenesis. Furthermore, to our best knowledge, this work demonstrates for the first time that the neural tube coverage by myotomes is driven by active cell movement, rather than passively by muscle growth. Consistent with this, in the absence of Zic1 activity, the myotome is unable to cover the neural tube at the hatching stage, despite increased dorsal myotome growth (Figures 1L and 2), supporting the involvement of the active cellular mechanism in this process. The model we propose for dorsal somite extension is shown in Figure 10.

Summary of dorsal somite extension in Wt and the Da mutant.

(A) Zic1 induces or maintains the expression of wnt11 during somite differentiation. This leads to a reduced proliferative activity of the dorsal DM and increases protrusion formation of the dorsal DM cells which are non-myogenic. Ultimately, the non-myogenic DM guides the epaxial myotomes dorsally where the myotomes form a gapless muscle layer covering the neural tube at the end of embryonic development. (B) In the Da mutant wnt11 expression is reduced in the dorsal somites. The dorsal DM cells show a high proliferative activity and reduced ability to form numerous large protrusions causing insufficient dorsal somite extension. This results in the incomplete coverage of the neural tube by the epaxial myotomes at the end of embryonic development.

In the present study, we described the characteristic behavior of dorsal DM cells and propose their guiding role for the epaxial myotome moving towards the top of the neural tube. During dorsal somite extension, DM cells at the tip of the dorsal somites form large, motile protrusions, which contain multiple bundles of filopodia dynamically branching out from the tip of the protrusions. These long protrusions could be beneficial for invading the restricted open space between the neural tube and the ectoderm. Additionally, previous studies showed that in migrating mesenchymal cells, the formation of lamellipodia is associated with higher migratory speed, whereas filopodia play an exploratory role and are associated with high directionality (Leithner et al., 2016; Innocenti, 2018). Large protrusions of wild-type DM cells consist of a lamellipodia-like core and multiple filopodia, which could account for fast dorsal somite extension with high accuracy. The detailed structure and function of the large protrusions needs to be further investigated in the future studies.

In zebrafish, hypaxial muscle precursors delaminate from the ventral tip of the DM and migrate collectively in a cell stream to the prospective pectoral fin bud region while the cells at the leading edge form long filopodia (Haines et al., 2004; Talbot et al., 2019). Thus, DM cells, regardless of whether dorsal or ventral, seem to have the potential to form protrusions and to exhibit migratory behavior. However, regulation of migratory behavior could be different as the migration of hypaxial muscle precursors is mediated by the receptor tyrosine kinase Met (Haines et al., 2004) which is not expressed in zebrafish dorsal DM cells.

At later stages of dorsal somite extension, we observed that DM cells delaminate from the tip of the dorsal somites and progressively occupy the space between the opposing dorsal somites. These mesenchymal DM cells actively form protrusions towards neighboring mesenchymal DM cells and DM cells of the somites, thus forming a dense cellular network on top of the neural tube. This could provide a communication platform for the opposing somites to meet at the right position, exactly on top of the neural tube. Indeed, ablation of mesenchymal DM cells resulted in transient opening of the gap between the somites, suggesting that these cells at least exert a tension to hold the positions of the dorsal tips of the left and right somite together during neural tube coverage. Interestingly, our lineage tracing experiment revealed that the dorsal DM cells at the tip of somites differentiated into cells of non-myotomal lineages including mural cells of blood vessels and mesenchymal cells in the dorsal fin fold. These observations are consistent with previous studies reporting their somitic origins in zebrafish and Xenopus (Garriock and Krieg, 2007; Lee et al., 2013; Ando et al., 2016). However, our data also showed that these DM cells do not appear to contribute to the myotome at the hatching stage. We thus propose that the neural tube coverage is driven by a specific population of dorsal DM cells, which do not contribute to myotomal cells during embryonic development.

The ability of the motile DM cells to guide the underlying myotome dorsally assures complete coverage of the neural tube by the epaxial myotomes. Considering the fact that DM cells are in close cell-cell contact with the underlying myotome, it is plausible that they are biomechanically coupled with myotomal cells, facilitating dorsal myotome extension. A similar mechanism has been investigated in mouse neural tube formation. During mouse neural tube closure, the surface ectoderm, overlying the neuroepithelium, forms cell protrusions towards the midline. Disruption of membrane ruffles, a form of lamellipodia, in the surface ectoderm results in incomplete fusion of the subjacent neuroepithelium (Rolo et al., 2016).

Our data suggest that in medaka dorsal somites Wnt11 exerts its effect through promotion of protrusion formation and down-regulation of cell proliferation in the dorsal DM. Regarding the protrusion formation, previous studies of migrating mesendodermal cells in the zebrafish mutant silberblick and neural crest cells in Xenopus embryos consistently reported that Wnt11 is involved in the oriented elongation and stabilization of protrusions (Ulrich et al., 2003; De Calisto et al., 2005; Matthews et al., 2008). Similarly, myocardial cells in the heart of Wnt11 mutant mice also form fewer protrusions (Zhou et al., 2007). Thus, regulation of cell protrusions is a conserved function of Wnt11, observed in various developmental processes. However, the relationship between Wnt11 and cell proliferation could be context-dependent, as it negatively regulates proliferation in mouse neonatal hearts (Touma, 2017), while it promotes cell proliferation in mouse intestinal epithelial cell culture (Ouko et al., 2004). Hence, the function of Wnt11 in medaka somites is unique in that it promotes tissue elongation by regulating a balance between proliferative and migrative activity.

By inhibiting the Wnt/Ca2+ signaling pathway we showed that Wnt11 probably acts through this non-canonical Wnt signaling pathway during dorsal somite extension. Likewise, previous studies in Xenopus have shown that Wnt11 acts through this pathway during cell migration into the dorsal fin fold. There it regulates epithelial–mesenchymal transition in a distinct dorsal somite cell population which, together with a population of neural crest cells, contribute to the mesenchyme of the dorsal fin fold (Garriock and Krieg, 2007). Furthermore, during convergent extension in vertebrate gastrulation, Wnt/Ca2+ signaling pathway can regulate cell adhesion by promoting actin polymerization (Choi and Han, 2002; Kohn and Moon, 2005). Whether Wnt11 functions cell-autonomously or not is context dependent (De Calisto et al., 2005; Garriock and Krieg, 2007; Gros et al., 2009). In our study, administration of exogenous Wnt11 protein rescued the protrusive behavior of the surrounding DM cells, suggesting that Wnt11 can function cell non-autonomously during dorsal somite extension.

Importantly, our RNA-seq and ChIPmentation analyses revealed that Zic1 has diverse downstream target genes including various developmental genes. This suggests that the dorsal myotome is established via pleiotropic actions of Zic1; Wnt11 may not be a sole factor for dorsal somite extension, although it was suggested to be crucial in the present study. The semaphorin-plexin pathway may play a role, since previous studies suggested, besides its implication in axon guidance and neural cell migration, a role in non-neural cell migration (Alto and Terman, 2017). Furthermore, cell migration and tissue deformation are often linked with the extracellular matrix (ECM), which provides guiding or restraining cues influencing cell movements. Previous studies showed that during mouse epaxial myotome development, ECM composition changes dynamically, which is tightly accompanied by epaxial muscle morphogenesis; while the laminin content in the ECM decreases, increasing tenascin and stable fibronectin contents potentially promotes the alignment of myofibers and their final organization (Deries et al., 2012). Furthermore, recent studies suggested that cells can remodel the surrounding ECM and thereby increase their own motility. For example, during zebrafish gastrulation, the metalloproteinase mmp14 is expressed by migrating endoderm cells and degrades laminin and fibronectin, components of the ECM (Hu et al., 2018). In this context, of particular importance is our identification of Adamts20, encoding a proteoglycanase, as a dorsal-high Zic1-target gene in the somites (Supplementary file 1). Since Adamts20 is known to play a pivotal role in embryonic melanoblast migration by remodeling the dermal ECM (Rao et al., 2003; Silver et al., 2008), Adamnts20 could thus facilitate migration of Wnt11-expressing DM cells through remodeling of the ECM. Further functional analysis of dorsal-high Zic1-target genes identified in somites will provide useful insight into the molecular network driving dorsal somite extension.

Finally, in vertebrates and especially in fish, body shape and muscle morphology are closely linked, since a majority of the body mass consist of muscular tissue. Previous studies in fish populations have shown that speciation and adaptation to a specific aquatic habitat are associated with a changes in body depth, a measurement of the trunk dorsoventral axis (Tobler, 2008; Elmer et al., 2010; Weese et al., 2012; Fruciano, 2016). In this context, the external appearance of the adult Da mutant is intriguing, in that it exhibits a teardrop body shape (typical for fish swimming in the middle layer like tuna), instead of a dorsally flattened one (typical for surface swimming fish like medaka). Since our study shows that the activity of wnt11 could influence the body shape by regulating cell proliferation and the behavior of muscle progenitor cells, Wnt11 could be one of the crucial factors in evolution and diversity of body shape in fish. Furthermore, the expression of zic1 (Ohtsuka et al., 2004; Sun Rhodes and Merzdorf, 2006; Houtmeyers et al., 2013) and wnt11 (Marcelle et al., 1997; Olivera-Martinez et al., 2004; Garriock et al., 2005; Matsui et al., 2005) are strongly conserved among vertebrates, and we hypothesize that Wnt11-mediated morphogenesis of the somites represents an evolutionarily conserved mechanism that acts across vertebrates.

Materials and methods

Key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Strain, strain background (Oryzias latipes)d-rRYamamoto, 1975Medaka Southern wild type population
Strain, strain background (O. latipes)Double anal fin (Da) mutantOhtsuka et al., 2004
Strain, strain background (O. latipes)Tg(zic1:GFP,zic4:DsRed)Kawanishi et al., 2013
Strain, strain background (O. latipes)Tg(zic1:zic1-Myc,zic4:DsRed);Dathis paperBAC construct zic1:zic1-Myc/zic4:DsRed was introduced into the Da mutant
AntibodyAnti-Digoxigenin-AP Fab fragments, peroxidase-conjugated (sheep polyclonal)Roche (Germany)11093274910IHC (1:2000)
AntibodyAnti-GFP (rabbit polyclonal)Clontech (Mountain View, California, USA)632592, RRID: 2336883IHC (1:500)
AntibodyAnti-mouse Alexa 555 (goat polyclonal)Thermo Fisher (Waltham, Massachusetts, USA)A-21422, RRID: AB_2535844IHC (1:500)
AntibodyAnti-Pax3/7 (DP312) (mouse monoclonal)Davis et al., 2001IHC (1:100)
AntibodyAnti-PCNA (rabbit polyclonal)Abcam (Cambridge)ab18197, RRID:AB_444313IHC (1:200)
AntibodyAnti-pH3 (Ser10) (rabbit polyclonal)Milipore (Burlington, Massachusetts, USA)06–570,RRID: AB_310177IHC (1:200)
AntibodyAnti-rabbit Alexa 488 (donkey polyclonal)Life Technologies (Carlsbad, California, USA)A-11008, RRID: AB_143165IHC (1:500)
Recombinant DNA reagentgRNA expression vector (DR274)Addgene (Watertown, Massachusetts, USA)42,250T7 promoter for in vivo transcription
Recombinant DNA reagentpCRIITOPO-wnt11Kawanishi et al., 2013For ISH at 12 ssT7 promoter for RNA generation
Recombinant DNA reagentpMTB-AC-TagGFP2this paperT7 promoter for mRNA generation
Recombinant DNA reagentpGEM-Teasy(Wnt11_ISH)this paperFor ISH at 22 ssSp6 promoter for RNA generation
Recombinant DNA reagentpMTB-memCherryXiong et al., 2014T7 promoter for mRNA generation
Recombinant DNA reagentpMTB-memb-mTagBFP2Collins et al., 2018Vector with Tol2 sites
Recombinant DNA reagentpMTB-t7-alpha-bungarotoxin J4 lab stockAddgene (Watertown, Massachusetts, USA)69,542T7 promoter for mRNA generation
Recombinant DNA reagentpMTB-KikGRThis paperSp6 promoter for mRNA generation
Recombinant DNA reagentpSPORT6.1(zic1-ISH)This paperT7 promoter for RNA generation
Recombinant DNA reagentzic1:GFP/zic4:DsRed (BAC)Kawanishi et al., 2013Vector with I-SceI meganuclease sites
Recombinant DNA reagentzic1:zic1-Myc/zic4:DsRed (BAC)This paperVector with I-SceI meganuclease sites
Sequence-based reagentPCR forward primer to amplify sgRNA template for in vivo transcriptionLee et al., 2020AAAAGCACCGACTCGGTG
Sequence-based reagentPCR reverse primer to amplify sgRNA template for in vivo transcriptionThis paperGGTCAGGTATGATTTAAATGGTCAGT
Sequence-based reagentPCR forward primer to generate wnt11 ISH probe for 12 ssKawanishi et al., 2013CAAATGGCTAACACTGTCTCAAAC
Sequence-based reagentPCR reverse primer to generate wnt11 ISH probe for 12 ssKawanishi et al., 2013CTATTTGCAAACGTATCTCTCCAC
Sequence-based reagentPCR forward primer to generate wnt11 ISH probe for 22 ssThis paperCATGAAGAGCCGCTCTCACA
Sequence-based reagentPCR reverse primer to generate wnt11 ISH probe for 22 ssThis paperTCCCTGAGGTCTTGGAGTCC
Sequence-based reagentRT-PCR forward primer for gapdhThis paperTGGCCATCAATGACCCGTTC
Sequence-based reagentRT-PCR reverse primer for gapdhThis paperTAGTTTGCCTCCCTCAGCCT
Sequence-based reagentRT-PCR forward primer for wnt11This paperCGCTGGAACTGCTCCTCTAT
Sequence-based reagentRT-PCR reverse primer for wnt11This paperGCAGCGGACAAGGCATAAAC
Sequence-based reagentRT-PCR forward primer for zic1Nakamura et al., 2014AGCCCTTTCCGTGTCCGTTCC
Sequence-based reagentRT-PCR reverse primer for zic1Nakamura et al., 2014CCGACGTGTGGACGTGCATGT
Sequence-based reagentStandard controlGene Tools (Philomath, Oregon, US)MorpholinoCCTCTTACCTCAGTTACAATTTATA
Sequence-based reagentwnt11 MOThis paperMorpholinoCTTCATGATGGATGGAGGCTCCGGT
Sequence-based reagentwnt11 PhotoMOThis paperMorpholinoCGGAGCCTPCATCCATCATG
Sequence-based reagentOligo annealing wnt11_1 sgRNA template, forwardThis paperTAGGTTCTGTCGGGAGACTTTA
Sequence-based reagentOligo annealing wnt11_1 sgRNA template, reverseThis paperAAACTAAAGTCTCCCGACAGAA
Sequence-based reagentOligo annealing wnt11_2 sgRNA template, forwardThis paperTAGGCTGGATGTTTAACGGAG
Sequence-based reagentOligo annealing wnt11_2 sgRNA template, reverseThis paperAAACCTCCGTTAAACATCCAG
Peptide, recombinant proteinBsaI-HFNew England Biolabs (Ipswich, USA)R3733
Peptide, recombinant proteinEnGen Spy Cas9 NLSNew England Biolabs (Ipswich, USA)M0646T
Peptide, recombinant proteinPancreatinWako (Japan)163–00142
Peptide, recombinant proteinRecombinant Human Wnt11 protein (hrWnt11)R&D systems (Minneapolis, Minnesota, USA)6179-WN-010/CF
Peptide, recombinant proteinTrypsinNacalai Tesque (Japan)35555–54
Chemical compound, drug4-nitro blue tetrazolium chloride (NBT)Roche (Germany)11585029001
Chemical compound, drug5-bromo-4-chloro-3-indolyl phosphate (BCIP)Roche (Germany)11383221001
Chemical compound, drugIsogenNippon Gene (Japan)315–02504
Chemical compound, drugKN-93Wako (Japan)114–00731
Chemical compound, drugML141Sigma-Aldrich (St. Louis, Missouri, USA)SML0407
Chemical compound, drugDAPISigma-Aldrich (St. Louis, Missouri, USA)D9542
Chemical compound, drugMayer’s Hematoxylin SolutionWako (Japan)131–09665
Chemical compound, drugPropidium iodineLife Technologies (Carlsbad, California, USA)P3566
Chemical compound, drugRhodamine PhalloidinLife Technologies (Carlsbad, California, USA)R415
Commercial assay or kitHiScribe T7 Quick High Yield RNA Synthesis KitNew England Biolabs (Ipswich, USA)E2050S
Commercial assay or kitKAPA HiFi HotStart ReadyMixKAPA Biosystems (Wilmington, Massachusetts, USA)KK2601
Commercial assay or kitKAPA Stranded mRNA-seq KitKAPA Biosystems (Wilmington, Massachusetts, USA)KK8420
Commercial assay or kitMinElute KitQiagen (Germany)28,004
Commercial assay or kitNextera Index KitIllumina (San Diego, California, USA)FC-121–1011
Commercial assay or kitNextera Sample Preparation KitIllumina (San Diego, California, USA)FC-121–1030
Commercial assay or kitRNeasy Mini KitQiagen (Germany)74,104
Commercial assay or kitSuper Script III KitInvitrogen (Waltham, Massachusetts, USA)18080051
Commercial assay or kitTHUNDERBIRD SYBR qPCR MixToyobo (Japan)QPS-201
Software, algorithmAdobe IllustratorAdobe (Mountain View, California, USA)Version 25.2.2
Software, algorithmBurrows-Wheeler Aligner (BWA) mapping softwareLi and Durbin, 2009
Software, algorithmFluoRenderWan et al., 2009Version 2.19
Software, algorithmHOMERHeinz et al., 2010
Software, algorithmImagej/ FijiSchindelin et al., 2012
Software, algorithmMACS2Zhang et al., 2008Version 2.1.1.20160309
Software, algorithmSTARDobin et al., 2013
Software, algorithmTrimmomaticBolger et al., 2014Version 0.32

Animals and transgenic lines

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Fish were raised and maintained under standard conditions. All experimental procedures and animal care were performed according to the animal ethics committee of the University of Tokyo. Sex was randomly assigned to experimental groups. Medaka d-rR stain was used as wild type, the Da mutant used in this study was previously described (Ohtsuka et al., 2004). The pre-existing transgenic line Tg(zic1:GFP,zic4:DsRed) (Kawanishi et al., 2013) was used, and the transgenic line Tg(zic1:GFP,zic4:DsRed);Da was created by crossing Da mutants with Tg(zic1:GFP,zic4:DsRed). The transgenic line Tg(zic1:zic1-Myc,zic4:DsRed);Da was generated by modifying the BAC used to generate Tg(zic1:GFP,zic4:DsRed) (Kawanishi et al., 2013), by replacing the ORF of GFP with the ORF of zic1 containing a sequence for a Myc-tag fused to its C-terminus. To establish the transgenic line, the BAC(zic1:zic1-Myc,zic4:DsRed) was co-injected with I-SceI Meganuclease (NEB) into one-cell stage Da embryos, as previously described (Thermes, 2002).

Visualization of actin skeleton of protrusions

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The AC-TagGFP2 sequence from the Actin-Chromobody plasmid (TagGFP2) (Chromotek) was cloned into the pMTB vector for mRNA generation. To investigate the actin skeleton of protrusions, cells of Wt and Da embryos were mosaically labeled using Actin-Chromobody-GFP (AC-GFP) mRNA. Embryos were injected at one-cell stage with 152 ng/µl membrane-mCherry mRNA. At four-cell stage, one cell was injected with 184 ng/µl AC-GFP mRNA. Embryos were raised until the desired stage and in vivo imaging was performed.

Lineage tracing of mesenchymal DM cells

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Mosaic labeling of cells was achieved by co-injecting 20 ng/µl pMTB-memb-mTagBFP2 plasmid with Tol2 mRNA into one-cell stage embryos of the transgenic line Tg(zic1:GFP,zic4:DsRed). Embryos were raised until 5 dpf and labeled cells were continuously observed until 8 dpf. Observation of cells was performed according to “In vivo imaging and in vivo time-lapse imaging”.

Photoconversion of Dorsal DM Cells

Wild type embryos were injected with 200 ng/µl kikGR mRNA at one-cell stage. Single dorsal DM cells exhibiting protrusions at the dorsal tip of 10th and 20th somites at 3.5 dpf (st. 29) were selectively photoconverted with a 405 nm laser under an LSM 710 confocal microscope and imaged again at 9 dpf (hatching stage). During the experiment, 8 embryos were labeled at both 10th and 20th somites, and 1 embryo was labeled only at the 10th somite.

Morpholino injection

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wnt11 MO was injected at one-cell stage. We used 12.5 µM wnt11 MO after finding that injection of 25 µM wnt11 MO gives rise to gastrulation delay, leading to smaller head and trunk phenotypes at the onset of dorsal somite extension. Injection of 12.5 µM wnt11 MO mostly did not hamper gastrulation. We excluded any MO-injected embryos showing gastrulation defects in the analyses at later timepoints.

Photo-Morpholino injection

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wnt11 Sense-Photo-Morpholino (PhotoMO) and wnt11 antisense Morpholino were annealed in a ratio 2:1. Microinjection of 25 µM of the annealed oligonucleotides was performed at one-cell stage, and embryos were raised until 4 ss in the dark. Photo-cleavage was performed using the 10 x objective and the DAPI filter of a Keyence BZ-9000 Biorevo microscope (Keyence). Embryos were mounted, dorsal side facing up, in 1% Methylcellulose in a glass bottom dish (Wako) and illuminated for 30 min. After Photo-cleavage, embryos were dechorionated and raised until the desired stage for subsequent analysis.

Generation of sgRNAs

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The online tool CCTop (Stemmer et al., 2015) was used to design sgRNAs. sgRNAs were generated as previously described by Hwang et al., 2013. Oligonucleotides were annealed and ligated into DR274 vector (Addgene), which was previously linearized using BsaI-HF (New England Biolabs). Template for in vivo transcription of sgRNA was amplified by PCR. In vivo transcription was performed using the HiScribe T7 Quick High Yield RNA Synthesis Kit (New England Biolabs) and RNA was purified using the RNeasy Mini kit (Qiagen). Quantity and quality of sgRNA were analyzed using a NanoDrop and agarose gel electrophoresis. sgRNA cleaving activity was confirmed by an in vivo assay.

sgRNA descriptionTarget site [PAM]
wnt11_1TTCTGTCGGGAGACTTTA[TGG]
wnt11_2GCTGGATGTTTAACGGAG[TGG]

wnt11 sgRNA injection

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Microinjections were performed using 25 ng/µl wnt11_1 sgRNA, 25 ng/µl wnt11_2 sgRNA and 100 ng/µl Cas9-NLS (New England Biolabs) at one-cell stage. To analyze phenotypes, embryos were mounted in 1% methylcellulose and imaged using a Leica M165 FC stereo microscope.

Injection of human recombinant Wnt11 protein into Da mutant somite

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To immobilize embryos, embryos from the Tg(zic1:GFP,zic4:DsRed);Da transgenic line were injected with 25 ng/µl α-bungarotoxin mRNA at one-cell stage. At 18 ss embryos were mounted in 1% low melting agarose in 1 x Yamamoto’s Ringer Solution and oriented with the dorsal side facing upwards. Embryos were injected on top of the 10th somite with a mix containing Dextran Rhodamine (Thermo Fisher) and 1.7 ng hrWnt11 protein (R&D Systems) or BSA (Sigma-Aldrich) and raised to 24 ss, followed by in vivo imaging and analysis.

Laser ablation of mesenchymal DM cells

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Tg(zic1:GFP,zic4:DsRed) embryos were immobilized by injecting 25 ng/µl α-bungarotoxin mRNA at one-cell stage and raised until 5.5 dpf. Z-stacks of the 9th (control somite) and 10th somite were imaged before ablation using the 40 x objective of a Zeiss LSM 710 confocal microscope. Laser ablation protocol was adapted from Morsch et al., 2017 and Volpe et al., 2020. Prior to laser ablation the 40 x objective and additional the 2.2 x zoom were used to focus only on the ablation site, the mesenchymal DM cells between the 10th somites. The ablation site was chosen to prevent damage of dorsal somites and the neural tube. Laser ablation was perforemd using a 405 nm laser at 80% power for 10 min on ‘continous’ mode. The 9th and 10th somite were subject to in vivo imaging directly after laser ablation and every 8 h until 40 hpa.

KN-93 treatment

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Dechorionated 4 ss embryos were treated with 30 μM KN-93 (Wako) or DMSO (Sigma-Aldrich) in 1 x Yamamoto’s Ringer Solution at 28 °C, in the dark until 22 ss.

ML141 treatment

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Embryos at 4 ss were dechorionated and treated with 500 μM ML141 (Sigma-Aldrich) or DMSO in 1 x Yamamoto’s Ringer Solution at 28 °C, in the dark until 24 ss.

In vivo imaging and in vivo time-lapse Imaging

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To immobilize embryos for in vivo imaging, embryos were injected at one-cell stage with 25 ng/µl α-bungarotoxin mRNA (Swinburne et al., 2015; Lischik et al., 2019). Embryos were mounted in 1% Low melting agarose (Sigma-Aldrich) in 1 x Yamamoto’s Ringer Solution in a glass-bottomed petri dish (IWAKI) and oriented with the dorsal side facing down. Imaging was performed using a Zeiss LSM 710 confocal microscope system (Zeiss) equipped with an inverted stand and a Zeiss AXIO Observer Z1 and a T-PMT detector. The embryos were positioned with the 5th or 10th somite in the center and images were acquired using a 40 x water objective. For the in vivo time-lapse imaging, the 10th or 15th somite was positioned in the center, z-stacks were imaged in a 600 s interval for 10–15 hr. Image analysis was performed in Fiji using the “Image Stabilizer” Plugin, the FFT Bandpass filter and the “Draw_arrows” Plugin to draw customized arrows (Li, 2008; Daetwyler et al., 2020).

Whole mount in situ hybridization

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Whole mount in situ hybridization was performed as previously described with the following modifications (Takashima et al., 2007). Embryos were fixed in 4% PFA/1.5 x PTW at 4 °C, overnight. Hybridization was performed at 65 °C, overnight. Samples were treated with alkaline-phosphatide anti-DIG-AP Fab fragments (1:2000, Roche). Signals were developed using 4-nitro blue tetrazolium chloride (NBT, Roche) and 5-bromo-4-chloro-3-indolyl phosphate (BCIP, Roche).

Whole-mount immunohistochemistry

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Embryos were fixed in 4% PFA/PBS for 2 hr at room temperature or at 4 °C overnight. Samples were permeabilized with 0.5% TritonX-100 (Wako) in 1 x PBS for 1–2 hr and blocked in blocking solution (2% BSA (Sigma-Aldrich), 1% DMSO, 0.2% TritonX-100 in 1 x PBS) for 2–4 hr at room temperature. Samples were incubated with respective primary antibody diluted in blocking solution at 4 °C, overnight. After an additional 4 hr blocking step, samples were incubated with respective secondary antibodies diluted in blocking solution, at 4 °C, overnight. Samples were stored in 1 x PBS at 4 °C until imaging.

Vibratome sectioning

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Samples were mounted in 4% agarose in 1 x PBS. 40 μm or 200 µm sections were obtained by a Vibratome (Leica, Vibratome). The sections were mounted on a glass slide (Matsunami) in 60% glycerol (Merck, Wako) and stored at 4 °C until imaging. Images were acquired using the 40 x water objective of a Zeiss LSM 710 confocal microscope.

Histological sections

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Dechorionated embryos were fixed in Bouin’s solution overnight, followed by a gradual dehydration using ethanol. Samples were embedded in Technovit 7,100 (Heraeus Kulzer) and sectioned into 5–6 µm thick sections. Sections were stained with hematoxylin (Wako) and imaged using the 1.6 x objective of a Leica M165 FC fluorescent stereo microscope.

Image processing and statistical analysis

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Image processing was performed with the image processing software Fiji. The 3D-recreation of in vivo imaging date was created using FluoRender (Wan et al., 2009). Measurements of morphological features (distance between myotome tips, area of cross-section of dorsal somites, diameter of myofibers, distance between dorsal somite tip and the tip of neural tube, somite height) was performed by averaging the analysis of the feature from three consecutive Y planes. RStudio was used for the statistical analysis and representation of the data. In bar plots mean and error limits, defined by the standard deviation, are indicated. In box plots median first and third quantiles are indicated. Statistical significance was determined by un-paired t-tests, a p-value p < 0.05 was considered as significant. In the figure legends sample size (n) and number of individuals used in the experiment are stated. Sample sizes were not predetermined using statistical methods, but the sample sizes used are similar to those generally used in the field. To compare experimental groups, the allocation was performed randomly, without blinding.

RT-PCR of cDNA generated from embryonic tails

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To investigate the gene expression in tails of embryos, tails were dissected anterior from the first somite. Ten tails were pooled together and RNA was isolated using Isogen (Nippon Gene). RNA was purified using the RNeasy Mini kit (Qiagen) and reverse transcribed to cDNA using the Super Script III Kit (Invitrogen). RT-PCR was performed using the Thunderbird Sybr qPCR Mix (Toyobo) following manufacturer’s instructions and run in the Agilent Mx3000P qPCR System (Agilent). Normalization of relative quantities was performed against gapdh expression, followed by analysis with excel and RStudio.

Isolation of dorsal and ventral somite cells for ATAC-Seq and RNA-Seq

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Yolk and head region were removed from 22 ss Tg(zic1:GFP,zic4:DsRed) embryos, and remaining trunk-tail pieces were incubated with 10 mg/ml pancreatin (Wako) at room temperature for 5–10 min to loosen the adhesion between tissues while maintaining the integrity within each tissue. Epidermis and intestinal tissues were removed, and then somites were isolated from the neural tube. Collected somites were subsequently dissociated into individual cells in 0.5% (w/v) Trypsin (Nacalai Tesque) at 37 °C for 10 min, and the dissociation was stopped by adding the same volume of 15% (v/v) FBS / Leiboviz’s L-15 (Life Technologies). Dissociated cells were washed with PBS, and sorted into GFP-positive (dorsal) and negative (ventral) cells using FACSAria III (BD Biosciences). Dead cells were detected by Propidium iodide (Life Technologies) and removed.

RNA-Seq

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Total RNA was extracted from sorted somite cells using RNeasy Mini kit (Qiagen). mRNA was enriched by poly-A capture and mRNA-seq libraries were generated using KAPA Stranded mRNA-seq Kit (KAPA Biosystems). Libraries were generated from two biological replicates, and sequenced using the Illumina HiSeq 1,500 platform.

RNA-Seq data processing

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The sequenced reads were pre-processed to remove low-quality bases and adapter derived sequences using Trimmomatic v0.32 (Bolger et al., 2014), and aligned to the medaka reference genome (HdrR, ASM223467v1) by STAR (Dobin et al., 2013). Reads with mapping quality (MAPQ) larger than or equal to 20 were used for the further analyses.

ATAC-Seq

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ATAC-seq was performed as previously described (Buenrostro et al., 2013) with some modifications. Approximately 4000 sorted somite cells were used for each experiment. After washing with PBS, cells were resuspended in 500 μl cold lysis buffer (10 mM Tris-HCl pH 7.4, 10 mM NaCl, 3 mM MgCl2, 0.1% Igepal CA-630), centrifuged for 10 min at 500 g, supernatant was removed. Tagmentation reaction was performed as described previously (Buenrostro et al., 2013) with Nextera Sample Preparation Kit (Illumina). After tagmented DNA was purified using MinElute kit (Qiagen), two sequential PCRs were performed to enrich small DNA fragments. First, a 9-cycle PCR was performed using indexed primers from Nextera Index Kit (Illumina) and KAPA HiFi HotStart ReadyMix (KAPA Biosystems), amplified DNA was size selected for a size less than 500 bp using AMPure XP beads (Beckman Coulter). A second 7-cycle PCR was performed using the same primers as for the first PCR. PCR product was purified by AMPure XP beads. Libraries were generated from two biological replicates, and sequenced using the Illumina HiSeq 1,500 platform.

ChIPmentation

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Yolk and head region were removed from 22 ss Tg(zic1:zic1-Myc,zic4:DsRed);Da embryos, followed by an incubation with 10 mg/ml pancreatin (Wako) at room temperature for 5–10 min. Epidermis and intestinal tissues were removed and somites were isolated from the neural tube. ChIP was performed as previously described with the following modifications (Nakamura et al., 2014). Isolated somites were fixed with 1% formaldehyde for 8 min at room temperature then quenched by adding glycine (200 mM final) and incubating on ice for 5 min. After washing with PBS, cell pellets were stored at –80 ℃. Approximately 1.8 × 106 cells were thawed on ice, suspended in lysis buffer (50 mM Tris-HCl (pH 8.0), 10 mM EDTA, 1% SDS, 20 mM Na-butyrate, complete protease inhibitors, 1 mM PMSF) and sonicated 10 times using a Sonifier (Branson) at power 5. Chromatin lysates were collected by centrifugation and diluted 10-fold with RIPA ChIP buffer (10 mM Tris-HCl (pH 8.0), 140 mM NaCl,1 mM EDTA, 0.5 mM EGTA, 1% Triton X-100, 0.1% SDS, 0.2% sodium deoxycholate, 20 mM Na-butyrate, complete protease inhibitors, 1 mM PMSF) followed by an incubation with antibody/protein A Dynabeads (Invitrogen) complex at 4 °C, overnight, while rotating. Immunoprecipitated samples were washed three times with RIPA buffer (10 mM Tris-HCl (pH 8.0), 140 mM NaCl,1 mM EDTA, 0.5 mM EGTA, 1% Triton X-100, 0.1% SDS, 0.2% sodium deoxycholate) and once with TE buffer. After the washing steps 150 μl of Tris-HCl was added.

Library preparation for ChIPmentation was performed as previously described (Schmidl et al., 2015) with the following modifications. Twenty-four μl of Tagmentation reaction mix (10 mM Tris-HCl pH8.0, 5 mM MgCl2, 10%(v/v) N,N-dimethyl formamide) and 1 μl of Tagment DNA Enzyme from Nextera Sample Preparation Kit (Illumina) were added to the DNA-beads complex and incubated for 70 s at 37 °C. A total of 150 μl ice-cold RIPA buffer was added and incubated for 5 min on ice. The DNA-beads complex was washed with RIPA buffer and TE buffer, suspended in 50 μl lysis buffer and 3 μl of 5 M NaCl, and incubated at 65 °C, overnight. The sample was incubated for 2 hr with 2 μl of 20 mg/ml ProteinasK (Roche), and subjected to AMPure XP beads (Beckman Coulter) purification. The library was amplified by 18-cycle PCR using indexed primers from Nextera Index Kit (Illumina) and KAPA HiFi HotStart ReadyMix (KAPA Biosystems). For the input chromatin, tagmentation reaction was performed after DNA purification. Libraries were generated from two biological replicates, and sequenced using the Illumina HiSeq 1,500 platform.

ChIPmentation and ATAC-Seq data processing

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The sequenced reads were pre-processed to remove low-quality bases and adapter derived sequences using Trimmomatic v0.32 (Bolger et al., 2014) and aligned to the medaka reference genome (HdrR, ASM223467v1) by BWA (Li and Durbin, 2009). Reads with mapping quality (MAPQ) larger than or equal to 20 were used for the further analyses. MACS2 (version 2.1.1.20160309) (Zhang et al., 2008) was used to call peaks and generate signals per million reads tracks using following options; ChIPmentation: -g 600000000 -B --SPMR --keep-dup 2, ATAC-seq: --nomodel --extsize 200 --shift –100 g 600000000 -q 0.01 -B --SPMR. The input chromatin was used as control of ChIPmentation.

For ChIPmentation, peak regions called by two biological replicates were used as reliable peaks.

Motif analyses of Zic1 ChIPmentation peaks

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Motifs enriched at reliable ChIPmentation peaks were analyzed by findMotifsGenome command of HOMER (Heinz et al., 2010) using default parameters.

Identification of Zic1 target genes

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Differentially expressed genes were identified using DESeq2 (padj <0.01). Each reliable ChIP peak was associated to the nearest TSS, and the gene was defined as Zic1-target gene if the distance between the peak and the TSS was closer than 50 kb.

Gene ontology and pathway analyses

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The gene ontology enrichment analyses and pathway enrichment analyses were performed using the Gene Ontology Resource (Ashburner et al., 2000; Gene Ontology Consortium, 2021; Gene Ontology Consortium, 2021).

Data availability

Sequencing data generated in this study have been submitted to the DDBJ BioProject database under accession number PRJDB11712.

The following data sets were generated
    1. Helig AK
    2. Nakamura R
    3. Hashimoto Y
    (2022) DDBJ BioProject
    ID PRJDB11712. The function of transcription factor Zic1 in dorsalization of the medaka somite.

References

    1. Hollway G
    2. Currie P
    (2005) Vertebrate myotome development
    Birth Defects Research. Part C, Embryo Today 75:172–179.
    https://doi.org/10.1002/bdrc.20046
  1. Book
    1. Romer A
    2. Parsons TS
    (1986)
    The Vertebrate Body
    Saunders College Pub.
    1. Tombes RM
    2. Grant S
    3. Westin EH
    4. Krystal G
    (1995)
    G1 cell cycle arrest and apoptosis are induced in NIH 3T3 cells by KN-93, an inhibitor of CaMK-II (the multifunctional Ca2+/CaM kinase
    Cell Growth & Differentiation 6:1063–1070.
  2. Book
    1. Yamamoto T
    (1975)
    Medaka (Killifish): Biology and Strains
    Tokyo: Keigaku Publishing Company.

Decision letter

  1. Tatjana Piotrowski
    Reviewing Editor; Stowers Institute for Medical Research, United States
  2. Marianne E Bronner
    Senior Editor; California Institute of Technology, United States

Our editorial process produces two outputs: i) public reviews designed to be posted alongside the preprint for the benefit of readers; ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Decision letter after peer review:

Thank you for submitting your article "Zic1 advances epaxial myotome morphogenesis to cover the neural tube via Wnt11r" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Marianne Bronner as the Senior Editor. The reviewers have opted to remain anonymous.

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential revisions:

All three reviewers agree that the study addresses an interesting and underexplored question in developmental biology. The findings are interesting, especially the illustration of the dynamic behaviors of dorsal somitic cells, which form elaborate protrusions, delaminate from their parent somite, and bridge the gap between opposing epaxial myotomes. The reviewers would like the authors to address the minor technical issues outlined in the individual reviews if possible, as well as the following major critiques:

1) The reliance on photomorpholinos to downregulate wnt11r dampens the enthusiasm of the reviewers, as the judgement of its efficacy relies on circumstantial evidence that the phenotype resembles the zic1 phenotype and that exogenously supplied human wnt11 protein partially rescues the phenotype. The reviewers encourage the authors to generate a wnt11r Crispr mutant to validate the morpholino experiments.

2) The identification of a migratory population of dorsal somitic cells is an intriguing observation and the focus of this paper. The fate and function of these cells in epaxial myogenesis, however, is not clear from the current data set and is a gap in the paper. The function of these cells could possibly be explored through an examination of mutants and treatments that prevent their formation or migration.

Reviewer #1 (Recommendations for the authors):

Critique:

1) Wnt11r in zebrafish was renamed to wnt11f1 and is the ortholog to human wnt11 (see Postlethwait et al. Evolutionary origin and nomenclature of wnt11- family genes, 2019). The authors should check if medaka wnt11r is the ortholog to human wnt11 and should also be renamed.

2) In zebrafish neither wnt11 or wnt11r mutants die during gastrulation. The authors should generate mutants before concluding that they will not survive past gastrulation, especially since there is no good control for the efficacy of the knockdown of wnt11r by photomorpholinos.

3) Figure 5A: It is unclear to me why zic1:GFP and zic4DSRed fish were generated. Wouldn't the DSRed interfere with the detection of dead PI+ cells during FACS sorting?

4) Figure 5C: Please describe in the result section that dorsal and ventral somites were dissected and that the GFP- cells do not consist of cells from elsewhere in the body.

5) Discussion: there is no discussion on how wnt11r might instruct polarized cell protrusions. Does wnt11r need to be expressed in a gradient? Does it act non-cell autonomously or in a paracrine fashion? Is wnt11r an instructive cue or a permissive cue?

6) Throughout the figures it is hard to see which panels show mutants and which ones show wildtype embryos. Please label better, eg. on the left of the row of images.

Reviewer #2 (Recommendations for the authors):

Pigmentation (e.g., Fig1I) makes it difficult to visualize the margins of the epaxial myotomes.

The authors should consider complementing Ph3 labelling with BrdU/Edu labelling for cell birth-dating as a measure of proliferation.

While the medial movement of Zic1:GFP cells to bridge the gap between epaxial myotomes is an interesting observation, the authors do not provide clear evidence of the function of this population in epaxial myotome closure of the neural tube. What is the fate of these cells? Do they contribute to epaxial musculature?

In Figure 4, it is unclear what BFP labeling marks.

The authors provide quantification of the protrusions between WT and Da mutants. The authors should consider a similar quantification for the number of dorsal mesenchymal cells between Da and Wt mutants. Additionally, since the authors have the data already in hand, rendered cross sections might provide another way to clearly visualize differences in the epaxial myotome between WT and mutant fish.

The authors identified Wnt11r as a putative mediator of Zic function during epaxial myotome expansion and hypotheisized Wnt11r functions through the Wnt/ca2+ signalling pathway. To test this, they treated embryos with Kn-93. Is it known whether this drug affects off additional pathways? The authors should include a comment on the specificity of the drug.

The authors should further explain the controls used for ChIP-seq experiments.

It would be interesting to know if similar cellular protrusions are present in the ventral margins of hypaxial myotomes, and if the presence of protrusions differ between fin and non-fin levels?

The role of migratory somitic cells to form hypaxial muscles such as those of limb is well described. The identification of migratory somitic cells originating from the Pax3/7+ DM/ECL population in fish is a particularly intriguing observation that seems to raise questions about how programs for migratory cell formation are similar and different between epaxial and hypaxial embryonic contexts.

Line 757 typo "semaphoring" should be "semaphorin".

The author's ChipSeq data sets highlight the position of putative Wnt11r enhancer, and their ATACseq data suggests this region is differentially open in the dorsal somite versus the ventral somite. Given these intriguing data sets, the authors should consider using this region to generate an enhancer-reporter transgenic line to test if this enhancer drives dorsal expression.

Reviewer #3 (Recommendations for the authors):

The opening several paragraphs of the discussion include materials that are already covered in the results. It's good to give a synopsis and explain the model, but it would be nice to see this manuscript better connected with the larger literature. Help the readers understand why this work is important. A few potential connections are laid out in the following paragraphs.

I find it striking how similar the cell behaviors are in the epaxial myotome to behaviors shown in the hypaxial myotome of zebrafish. For instance, in both cases researchers have used long-protrusions as evidence of cellular guidance. It would be interesting to see discussion added comparing/contrasting behaviors on the extreme dorsal/ventral ends of differing somites. I will understand if you think this is off-topic, but I think a little discussion about how the epaxial findings related to other muscle groups could give context that broadens the impact of the study.

Even in the narrow literature, some citations are missing. For instance, a quick pubmed of 'zic1 epaxial' gives only three hits; one by the authors and two from another group – the latter two are not cited; these could easily be worked into introduction or discussion.

The paper could also benefit from the authors finding that Da mutants show increased dorsal muscle growth compared to the wild type, even though the mutant muscles do not enclose the dorsal surface. This finding seems like it further supports the author's claim that dorsal closure is not controlled merely by physical constraint; if the authors agree, they could mention this in the discussion.

Presentation issues:

The time-lapse videos could be strengthened by additional annotation of what the reader is supposed to be looking at. The one little arrow in videos 1 and 2 helps some, but it is not enough when there are complex movements amongst a sea of cells and no annotations are provided for videos 3-5, even though arrowheads are mentioned in-text for videos 3 and 4.

I needed to modify videos 3-5 before they played on my new Mac, though they did on a somewhat older Mac. Please double-check the formatting and make sure they are cross compatible with newer systems. The lack of arrowheads also suggests that an incorrect version may have been uploaded.

The animals in Video 3 (especially) and 5 do not hold still, making it difficult to follow what's happening. If feasible amid covid restrictions, it would be nice to see these replaced with time-lapse of immobilized fish.

It is difficult to see protrusions in figure 8b. This panel would benefit from separation of Rhodamine/GFP channels (in addition to the merge) and/or an accompanying illustration.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "Zic1 advances epaxial myotome morphogenesis to cover the neural tube via Wnt11" for further consideration by eLife. Your revised article has been evaluated by Marianne Bronner (Senior Editor) and a Reviewing Editor.

The manuscript has been significantly improved but there are some remaining issues that need to be addressed, as outlined below:

As noted in the original reviews, the reviewers find the results reported in the manuscript interesting and the data of high quality. The revised manuscript contains new experimental data and the majority of the reviewers' prior concerns are addressed.

The reviewers agree that the requested wnt11 mutant analysis is indeed not informative as the embryos possess convergent extension defects. Even though the wnt11 morpholino experiments possess caveats, their phenotype correlates with the results of several other experiments, suggesting that the morpholinos work as expected. However, the text describing the results should reflect that these are morpholino experiments. Eg., line 337 could be changed to saying that the MO 'supports an important role'.

One major concern with the prior submission was that the fate and function of the dorsal somitic cells bridging opposing myotomes were not described. Here the authors present new data from pharmacological perturbation experiments and targeted laser ablation experiments. These data improve the manuscript and provide evidence that somitic mesenchymal cells have a functional role in myotome extension around the neural tube.

Unfortunately, a more detailed fate analysis of these cells is still missing. A fascinating finding of the paper is the time-lapse analysis of the cells crawling around on top of the neural tube. But it is not entirely clear what those cells become – are they becoming the fin fold or are they becoming muscle? They do not look too different from the fin mesenchyme cells Tom Carney looked at in a paper a few years back, and their appearance does look different from some zebrafish epaxial muscle-forming cells. Since the authors' main conclusion is about muscle, and their main analysis is of cells that we are not sure are becoming muscle, the authors should perform a more conclusive fate analysis.

So far, the fate of somitic mesenchyme cells is only partially addressed through use of mosaic pMTB-membmTagBFP2 expression. The authors extend their previous analysis to argue DM cells contribute to the median fin-fold. Did the authors examine only those somites where mosaic BFP was restricted to the DM and not present in other somitic tissues? This would be important for interpretation of results.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "Zic1 advances epaxial myotome morphogenesis to cover the neural tube via Wnt11" for further consideration by eLife. Your revised article has been evaluated by Marianne Bronner (Senior Editor) and a Reviewing Editor.

The manuscript has been improved but there are some remaining issues that need to be addressed, as outlined below:

The authors further improved the manuscript by adding fate mapping data of the dorsal DM cells. The fate mapping shows that the dorsally extending cells of interest contribute to the fin-fold mesenchyme and appear to surround blood vessels, however no labeled cells contribute to the myotome itself.

1) These new findings need to be better represented throughout the manuscript. For example, the abstract still reads:

'In wild type, dorsal dermomyotome (DM) cells, progenitors of myotomal cells, reduce their proliferative activity after somitogenesis and subsequently form unique large protrusions extending dorsally, guiding the epaxial myotome dorsally'

Please clarify that dorsal dermomyotome cells are not myotomal progenitors in this context.

2) Given the new fate mapping data, please also make the modified model more explicit throughout the abstract, intro and discussion: that a non-myogenic lineage of pioneer cells non-cell-autonomously guides dorsal myotome growth.

3) Could the authors confirm that the fate mapping experiments consist of 17 individual cells in 17 different embryos?

4) Title: 'Zic1 advances epaxial myotome morphogenesis to cover the neural tube via Wnt11'. Please consider rephrasing the title, which currently does not convey that wnt11 acts on DM cells, which then guide the epaxial myotome. The title suggests that wnt11 acts on epaxial myotome cells directly.

Line 571, 1371: CRISPR-mediated knock-out of wnt11. Please, rephrase knock-out as the analyzed embryos were not stable mutants. F0 CRISPR causes a hodgepodge of mutations and WT cells.

Please soften the language regarding the morpholino experiments as initially requested by the reviewers.

https://doi.org/10.7554/eLife.71845.sa1

Author response

Essential revisions:

All three reviewers agree that the study addresses an interesting and underexplored question in developmental biology. The findings are interesting, especially the illustration of the dynamic behaviors of dorsal somitic cells, which form elaborate protrusions, delaminate from their parent somite, and bridge the gap between opposing epaxial myotomes. The reviewers would like the authors to address the minor technical issues outlined in the individual reviews if possible, as well as the following major critiques:

1) The reliance on photomorpholinos to downregulate wnt11r dampens the enthusiasm of the reviewers, as the judgement of its efficacy relies on circumstantial evidence that the phenotype resembles the zic1 phenotype and that exogenously supplied human wnt11 protein partially rescues the phenotype. The reviewers encourage the authors to generate a wnt11r Crispr mutant to validate the morpholino experiments.

Following the reviewers’ suggestion, we designed gRNAs against wnt11 (we renamed the gene wnt11 from wnt11r after a suggestion by Reviewer 1) and generated wnt11 crispant embryos (Figure 7 —figure supplement 2A). We found out that development in the crispants were severely affected with a smaller head and truncated body axis at the somitogenesis stage (Figure 7 —figure supplement 2E-F’, arrowheads). This phenotype is known to be secondarily caused by impaired gastrulation movement, and indeed we observed gastrulation defects in the crispants (Figure 7 —figure supplement 2C, D). Because of these defects, we were unable to assess the myotome development in the wnt11 mutant. However, these phenotypes (impaired gastrulation movement and malformed trunk) were consistent with our observation in the wnt11 morphants when a high dose of wnt11 MO was injected, further supporting that the wnt11 MO inhibits the wnt11 gene specifically.

In the revised manuscript, we added description on the wnt11 cripant embryos to justify the MO experiments (Line 305).

“Injection of gRNAs against wnt11 to knock out the gene revealed that the resultant F0 embryos displayed delayed epiboly movement and subsequent morphological defects, including shorter body axis and impaired trunk development (Figure 7 —figure supplement 2), implying the difficulty of assessment of the wnt11 function at later stages using the wnt11 mutant.”

2) The identification of a migratory population of dorsal somitic cells is an intriguing observation and the focus of this paper. The fate and function of these cells in epaxial myogenesis, however, is not clear from the current data set and is a gap in the paper. The function of these cells could possibly be explored through an examination of mutants and treatments that prevent their formation or migration.

We absolutely agree with the suggestion that the function and fate of the dorsal DM and mesenchymal cells are important for the significance of this study. We tested their roles with two additional experiments.

1) At the onset of dorsal somite extension, we inhibited filopodia formation of large protrusions of dorsal DM cells with ML141, a specific inhibitor against Rac1/Cdc42 (Figure 3L, M), and found that the treatment disturbed the dorsal extension of myotomes (Figure 3N). This result suggests that the dorsal DM cells promote the dorsal somite extension by utilizing the large protrusions.

2) We also ablated the mesenchymal DM cells arising at later stages of dorsal somite extension, using a UV laser and revealed that the ablation transiently widened the gap between the left and right myotomes and delayed the dorsal somite extension (Figure 4J-L). We concluded that the mesenchymal DM cells exert a force promoting the advancement of left and right myotomes to cover the neural tube. Thus, DM cells seem to be playing a crucial role throughout dorsal somite extension.

We further investigated the fate of the dorsal DM cells by sparsely labeling those cells with BFP plasmid injection. We found that some cell eventually differentiated into myotome fibers while some became mesenchymal cells in the dorsal finfold (Figure 4 —figure supplement 3C). This result implies that the mesenchymal cells have multiple potentials to become at least myotomal and dermal cells.

We included the description on the function of the dorsal DM and mesenchymal cells in the Results section as follows:

1 (Line 172):

“To investigate the role of large protrusions during the onset of dorsal somite extension, we inhibited filopodia formation using ML141 (Figure 3L, M). ML141 specifically inhibits Cdc42/Rac1 GTPases, which are critical for filopodia formation (Hong et al., 2013; Fantin et al., 2015). Intriguingly, dorsal somites of embryos treated with ML141 extended significantly less dorsally compared to control embryos treated with DMSO (Figure 3N).”

2 (Line 208):

“To examine the function of mesenchymal DM cells, we ablated these cells between the 10th somites of 5.5 dpf Tg(zic1:GFP) embryos with a UV laser (Figure 4J-K’, Figure 4 —figure supplement 3A-B’). Intriguingly, the distance between the left and the right tips of the 10th somites increased after ablating the mesenchymal DM cells while the neighboring 9th somites continued to shorten the gap (Figure 4L). This suggests that mesenchymal DM cells hold the left and the right somite together to promote the dorsal somite extension at this late phase of myotome development. After 16 hours post ablation, dorsal extension of the 10th somites eventually resumed as the mesenchymal DM cells regenerated at the ablation site (Figure 4L).”

Regarding the fate of DM cells, we added a sentence “Interestingly, besides the differentiation into myotome cells, our preliminary data suggest that DM cells also differentiate into mesenchymal fin fold cells, aligning with previous observations in Xenopus (Garriock and Krieg, 2007). “ to the Discussion section (Line 423).

Reviewer #1 (Recommendations for the authors):

Critique:

1) Wnt11r in zebrafish was renamed to wnt11f1 and is the ortholog to human wnt11 (see Postlethwait et al. Evolutionary origin and nomenclature of wnt11- family genes, 2019). The authors should check if medaka wnt11r is the ortholog to human wnt11 and should also be renamed.

We thank the reviewer for the important suggestion. We examined the sequence homology and synteny of medaka wnt11r and confirmed that it is the only orthologue of human WNT11 (there was no wnt11f1 found in the medaka genome). We rewrote it as wnt11 throughout the manuscript.

2) In zebrafish neither wnt11 or wnt11r mutants die during gastrulation. The authors should generate mutants before concluding that they will not survive past gastrulation, especially since there is no good control for the efficacy of the knockdown of wnt11r by photomorpholinos.

Following the reviewer’s comment, we generated wnt11 crispant embryos and found that they have gastrulation defects which lead to impaired trunk development (Figure 7 —figure supplement 2). We also confirmed that the crispant phenotypes were identical to the phenotypes of wnt11 morphants after injected with a high dose of wnt11 MO. We thus think that we can evaluate the wnt11 function using the MO and PhotoMO.

We found that the medaka genome has only one wnt11 while there was (no wnt11f1 or a homologue of zebrafish silberblick). We reason that the severe phenotype observed in medaka wnt11 crispant embryos is probably due to the lack of paralogues (wnt11f1).

In the revised manuscript, we added description on the wnt11 cripant embryos to justify the MO experiments (Line 305).

“Injection of gRNAs against wnt11 to knock out the gene revealed that the resultant F0 embryos displayed delayed epiboly movement and subsequent morphological defects, including shorter body axis and impaired trunk development (Figure 7 —figure supplement 2), implying the difficulty of assessment of the wnt11 function at later stages using the wnt11 mutant.”

3) Figure 5A: It is unclear to me why zic1:GFP and zic4DSRed fish were generated. Wouldn't the DSRed interfere with the detection of dead PI+ cells during FACS sorting?

We apologize for the misleading figure. We used the zic1:GFP;zic4:DsRed transgenic line that had been used in the previous chapters. As described in Line 143, DsRed fluorescence in somitic cells is very faint due to the weak activity of the zic4 promoter. Thus the weak DsRed fluorescence did not interfere with the π staining. To avoid confusion, we removed the picture of BAC construct in Figure 5A.

4) Figure 5C: Please describe in the result section that dorsal and ventral somites were dissected and that the GFP- cells do not consist of cells from elsewhere in the body.

We thank the reviewer for the suggestion. We exclusively isolated somites after removing the rest of the embryos before FACS sorting. Thus GFP+ and – cells in the analyses are all derived from somites. Following the reviewer’s suggestion, we clarified the description on the somite dissection procedure in the Results section (Line 227), methods section (Line 665) and in Figure 5A,B.

5) Discussion: there is no discussion on how wnt11r might instruct polarized cell protrusions. Does wnt11r need to be expressed in a gradient? Does it act non-cell autonomously or in a paracrine fashion? Is wnt11r an instructive cue or a permissive cue?

According to previous studies, the cell autonomy of Wnt11 function is context dependent. In Xenopus finfold mesenchymal cells, Wnt11 was suggested to act in a cell-autonomous manner. Meanwhile, other studies have proposed a cell non-autonomous role of Wnt11 in Xenopus cranial neural crest migration and chick muscle fiber orientation (De Calisto et al. Development 2005; Gros et al. Nature 2009). In our study, administration of exogenous Wnt11 protein rescued the protrusive behavior of the surrounding DM cells, suggesting that Wnt11 can function cell non-autonomously during dorsal somite extension. Since the dorsal DM cells gradually migrate out of the wnt11-positive dorsal somite region during the process, Wnt11 may not work as an attractive (instructive) cue, but likely works as a permissive cue for dorsal migration.

However, our current data do not clearly tell whether Wnt11 functions cell-autonomously or non-cell-autonomously. We thus only added a few sentences to the discussion part Line 457 as follows:

“Whether Wnt11 functions cell-autonomously or not is context dependent (Garriock and Krieg, 2007; De Calisto et al. 2005; Gros et al. 2009). In our study, administration of exogenous Wnt11 protein rescued the protrusive behavior of the surrounding DM cells, suggesting that Wnt11 can function cell non-autonomously during dorsal somite extension.”

6) Throughout the figures it is hard to see which panels show mutants and which ones show wildtype embryos. Please label better, eg. on the left of the row of images.

We appreciate the reviewer’s comment. We added labels on the left of the images in most figures.

Reviewer #2 (Recommendations for the authors):

Pigmentation (e.g., Fig1I) makes it difficult to visualize the margins of the epaxial myotomes.

In our previous studies we have used the Da mutant strain derived from a natural population which has melanophores. We used the same strain in the current study for consistency of all data. Because the contour of the myotomes was sometimes difficult to see in a single z plane of confocal images as the reviewer pointed out, we carefully examined the z-stack images to determine the precise position of the boundaries. We explain this in the Figure legend, “the contour of the myotomes was drawn based on the Z-stack images of the dorsal myotomes to avoid ambiguity caused by melanophores.“

The authors should consider complementing Ph3 labelling with BrdU/Edu labelling for cell birth-dating as a measure of proliferation.

We tried EdU labelling, but unfortunately it did not work well for medaka embryos under our experimental condition. We thus performed an alternative experiment using immunohistochemistry against another proliferation marker, PCNA, to confirm the results of pH3 staining. In the Results section, we added the sentences accordingly (Line 126):

“Immunohistochemistry against another proliferation marker PCNA confirmed these findings (Figure 2 —figure supplement 1A-C).”

While the medial movement of Zic1:GFP cells to bridge the gap between epaxial myotomes is an interesting observation, the authors do not provide clear evidence of the function of this population in epaxial myotome closure of the neural tube. What is the fate of these cells? Do they contribute to epaxial musculature?

We agree with the reviewer’s suggestion and experimentally investigated the function and fate of the dorsal DM and mesenchymal cells. As described in detail in our response to Essential Revision 2 above, we have changed the manuscript by incorporating the new results (function – Lines 172, 208; fate – Line 423).

In Figure 4, it is unclear what BFP labeling marks.

We have revised Figure 4A-F to indicate more clearly the BFP-labelled somites and dorsally moving cells derived from them.

The authors provide quantification of the protrusions between WT and Da mutants. The authors should consider a similar quantification for the number of dorsal mesenchymal cells between Da and Wt mutants. Additionally, since the authors have the data already in hand, rendered cross sections might provide another way to clearly visualize differences in the epaxial myotome between WT and mutant fish.

We thank the reviewer for the important suggestions. We do agree that the difference between Wt and Da mutant mesenchymal DM cells can be clarified by providing cross-sections. In the revised manuscript, we thus added cross-sections to Figure 4A-F.

We also quantified the number of mesenchymal DM cells by analyzing the cross-sections of Tg(zic1:GFP) and Tg(zic1:GFP);Da embryos and incorporated the results into the main text (Figure 4 —figure supplement 1L, left graph). We found that there is a trend towards more mesenchymal cells accumulating between the somites of embryos in Wt background, but the difference was not statistically significant. We thus added a phrase “while the number of mesenchymal cells was not largely affected“ to the Results section (Line 206).

The authors identified Wnt11r as a putative mediator of Zic function during epaxial myotome expansion and hypotheisized Wnt11r functions through the Wnt/ca2+ signalling pathway. To test this, they treated embryos with Kn-93. Is it known whether this drug affects off additional pathways? The authors should include a comment on the specificity of the drug.

The specificity of KN-93 inhibition on CaMKII was previously validated (Sumi et al. Biochem Biophys Res Commun. 1991), and the drug has been widely used for inhibition of CaMKII in various studies (Tombes et al., 1995; Wu and Cline, 1998; Garriock and Krieg, 2007; Rothschild et al., 2013). We clarified the specificity of the drug in the main text (Line 355).

The authors should further explain the controls used for ChIP-seq experiments.

We used input chromatin for the control of ChIPmentation. The input chromatin of the ChIP was tagmentated after DNA purification step. This input chromatin was used for calling peaks by MACS2 software. We modified the methods to include the description on the control (Line 745).

It would be interesting to know if similar cellular protrusions are present in the ventral margins of hypaxial myotomes, and if the presence of protrusions differ between fin and non-fin levels?

We thank the reviewer for raising these intriguing questions. For technical reasons, it is hard to observe the extension procedure of the hypaxial myotomes. We speculate that the ventral-most DM cells also exhibit protrusions while they are less active than in dorsal DM cells, since the hypaxial somites show similar characteristics to the epaxial ones in the Da mutant embryos. The left and right myotomes cover the neural tube along the entire anterior-posterior axis including the dorsal fin level. It is thus likely that the protrusive activity of the dorsal DM cells can be observed regardless of the presence of fins.

The role of migratory somitic cells to form hypaxial muscles such as those of limb is well described. The identification of migratory somitic cells originating from the Pax3/7+ DM/ECL population in fish is a particularly intriguing observation that seems to raise questions about how programs for migratory cell formation are similar and different between epaxial and hypaxial embryonic contexts.

We agree that the similarity of the migratory behavior in the dorsal and ventral/hypaxial muscle precursor cells derived from DM would be an interesting topic. In the discussion part, we added the following sentences to compare the two systems in detail (Line 408):

“In zebrafish, hypaxial muscle precursors delaminate from the ventral tip of the DM and migrate collectively in a cell stream to the prospective pectoral fin bud region while the cells at the leading edge form long filopodia (Haines et al., 2004; Talbot et al., 2019). Thus, DM cells, regardless of whether dorsal or ventral, seem to have the potential to form protrusions and to exhibit migratory behavior. However, regulation of migratory behavior could be different as the migration of hypaxial muscle precursors is mediated by the receptor tyrosine kinase Met (Haines et al., 2004) which is not expressed in zebrafish dorsal DM cells.”

Line 757 typo "semaphoring" should be "semaphorin".

We apologize for the typo. We have corrected the spelling of semaphorin.

The author's ChipSeq data sets highlight the position of putative Wnt11r enhancer, and their ATACseq data suggests this region is differentially open in the dorsal somite versus the ventral somite. Given these intriguing data sets, the authors should consider using this region to generate an enhancer-reporter transgenic line to test if this enhancer drives dorsal expression.

We appreciate the reviewer’s helpful suggestion. To prove that the region acts as a wnt11 enhancer, it will be necessary to establish the enhancer-reporter line. However, we think that the enhancer analysis of the wnt11 gene is beyond the scope of this study. We therefore did not perform experiments to address this issue. In this regard, the use of the word “enhancer” may not be appropriate, and we therefore rewrote the Results section as follows (Line 291):

“These sites were more accessible in dorsal somites than in ventral somites, suggesting that Zic1 regulates wnt11 directly (Figure 6C).”

Reviewer #3 (Recommendations for the authors):

The opening several paragraphs of the discussion include materials that are already covered in the results. It's good to give a synopsis and explain the model, but it would be nice to see this manuscript better connected with the larger literature. Help the readers understand why this work is important. A few potential connections are laid out in the following paragraphs.

We thank the reviewer for the valuable advice. We shortened the first paragraphs in the discussion and added the following sentence to clarify the significance of our work (Line 388).

“To our knowledge, our work demonstrates for the first time that neural tube coverage by myotomes is driven by active cell movement, rather than being passively achieved by muscle growth.”

I find it striking how similar the cell behaviors are in the epaxial myotome to behaviors shown in the hypaxial myotome of zebrafish. For instance, in both cases researchers have used long-protrusions as evidence of cellular guidance. It would be interesting to see discussion added comparing/contrasting behaviors on the extreme dorsal/ventral ends of differing somites. I will understand if you think this is off-topic, but I think a little discussion about how the epaxial findings related to other muscle groups could give context that broadens the impact of the study.

We appreciate the reviewer’s suggestion. This interesting point was also raised by Reviewer 2, and we have incorporated the comparison between the dorsal and the ventral/hypaxial muscle precursor cells into the Discussion section as follows (Line 408):

“In zebrafish, hypaxial muscle precursors delaminate from the ventral tip of DM and migrate to the prospective pectoral fin bud region, extending filopodial protrusions (Haines et al. 2004; Talbot et al. 2019). DM cells, regardless of whether dorsal or ventral, could thus generally have potential to exhibit migratory behavior with active protrusions. However, regulation of migratory behavior could be different as the migration of hypaxial muscle precursors is mediated by the receptor tyrosine kinase Met (Haines et al. 2004) while it is not expressed in zebrafish dorsal DM cells. Furthermore, hypaxial muscle precursors form tightly packed cell streams migrating out from the somite, whereas dorsal DM cells migrate individually.”

Even in the narrow literature, some citations are missing. For instance, a quick pubmed of 'zic1 epaxial' gives only three hits; one by the authors and two from another group – the latter two are not cited; these could easily be worked into introduction or discussion.

We are sorry for the insufficient citations. We added several more papers referencing the functions of zic1/zic4 during somite development and specifically during myotome development in fish (Lines 40, 49).

The paper could also benefit from the authors finding that Da mutants show increased dorsal muscle growth compared to the wild type, even though the mutant muscles do not enclose the dorsal surface. This finding seems like it further supports the author's claim that dorsal closure is not controlled merely by physical constraint; if the authors agree, they could mention this in the discussion.

We fully agree with the reviewer’s opinion and incorporated this discussion in the Results section to justify our analyses of cellular behavior, as well as in the Discussion section.

(Line 136)

“In the Da mutant, the myotome is unable to cover the neural tube despite increased dorsal myotome growth at the hatching stage (Figure 1L, Figure 2). This suggest that additionally to physical extension an active process might support the dorsal movement of somites.”

(Line 391):

“Consistent with this, in the absence of Zic1 activity, the myotome is unable to cover the neural tube at the hatching stage despite increased dorsal myotome growth (Figure 1L, Figure 2), supporting the involvement of the active cellular mechanism in this process.”

Presentation issues:

The time-lapse videos could be strengthened by additional annotation of what the reader is supposed to be looking at. The one little arrow in videos 1 and 2 helps some, but it is not enough when there are complex movements amongst a sea of cells and no annotations are provided for videos 3-5, even though arrowheads are mentioned in-text for videos 3 and 4.

We apologize for the confusion. The legends of videos 3 and 4 in the result part do belong to the videos 1 and 2. There must have happened a mix-up of the video files. We reviewed the videos and legends in the main text again and uploaded the videos correctly.

I needed to modify videos 3-5 before they played on my new Mac, though they did on a somewhat older Mac. Please double-check the formatting and make sure they are cross compatible with newer systems. The lack of arrowheads also suggests that an incorrect version may have been uploaded.

We thank the reviewer for pointing out this issue. We modified the format of the videos and verified that they can be now played on new and old Mac computers, as well as on Windows computers. The lack of arrowheads in the videos 3 and 4 was due to the mishandling of annotations of the files (as explained in the comment above) and has been verified too.

The animals in Video 3 (especially) and 5 do not hold still, making it difficult to follow what's happening. If feasible amid covid restrictions, it would be nice to see these replaced with time-lapse of immobilized fish.

We appreciate the reviewer’s comment. We tried to fix the position of the embryos but we failed to do so completely for the following technical reasons. The movement of the medaka embryos in the videos consists of three factors: muscle contraction, yolk contraction and body axis growth. We successfully inhibited the muscle movement by injecting bungarotoxin mRNA which blocks nicotinic acetylcholine receptors (Swinburne et al. PLoS One 2015; Lischik et al. PLoS One 2019). However, the yolk contraction movement, which is unique to the early stages (from epiboly to mid-somitogenesis stages) of medaka embryos and is not observed in zebrafish, was difficult to be completely inhibited. The slight twitching of the embryos in the movies are due to this yolk movement. Furthermore, the movement to the left in videos 3 and 4 is due to the rapid body axis growth of the embryos during that stage, although we minimized the growth effect by registering the images by ImageJ.

It is difficult to see protrusions in figure 8b. This panel would benefit from separation of Rhodamine/GFP channels (in addition to the merge) and/or an accompanying illustration.

We fully agree with the reviewer and separated the channels in Figure 8B.

We also reorganized the formality of every figure legend to enhance the readability of them in the revised manuscript but did not highlight the changes using the Word proofreading function. Other modifications are highlighted.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

The manuscript has been significantly improved but there are some remaining issues that need to be addressed, as outlined below:

As noted in the original reviews, the reviewers find the results reported in the manuscript interesting and the data of high quality. The revised manuscript contains new experimental data and the majority of the reviewers' prior concerns are addressed.

The reviewers agree that the requested wnt11 mutant analysis is indeed not informative as the embryos possess convergent extension defects. Even though the wnt11 morpholino experiments possess caveats, their phenotype correlates with the results of several other experiments, suggesting that the morpholinos work as expected. However, the text describing the results should reflect that these are morpholino experiments. Eg., line 337 could be changed to saying that the MO 'supports an important role'.

One major concern with the prior submission was that the fate and function of the dorsal somitic cells bridging opposing myotomes were not described. Here the authors present new data from pharmacological perturbation experiments and targeted laser ablation experiments. These data improve the manuscript and provide evidence that somitic mesenchymal cells have a functional role in myotome extension around the neural tube.

Unfortunately, a more detailed fate analysis of these cells is still missing. A fascinating finding of the paper is the time-lapse analysis of the cells crawling around on top of the neural tube. But it is not entirely clear what those cells become – are they becoming the fin fold or are they becoming muscle? They do not look too different from the fin mesenchyme cells Tom Carney looked at in a paper a few years back, and their appearance does look different from some zebrafish epaxial muscle-forming cells. Since the authors' main conclusion is about muscle, and their main analysis is of cells that we are not sure are becoming muscle, the authors should perform a more conclusive fate analysis.

So far, the fate of somitic mesenchyme cells is only partially addressed through use of mosaic pMTB-membmTagBFP2 expression. The authors extend their previous analysis to argue DM cells contribute to the median fin-fold. Did the authors examine only those somites where mosaic BFP was restricted to the DM and not present in other somitic tissues? This would be important for interpretation of results.

We agree with the reviewers and indeed could not exclude the possibility that other somitic cells besides the dorsal DM cells were also labeled during our lineage tracing experiment using the BFP plasmid. To tackle this issue, we set up another experiment utilizing KikGR (photoconvertible protein)-mediated photoconversion, which enables spatiotemporally controlled labeling of dorsal DM cells at a single-cell level. We injected kikGR mRNA into 1-cell stage embryos and photoconverted single dorsal DM cells specifically with a UV laser. Specific labeling of the dorsal DM cells was confirmed by visual inspection of z-stack images using confocal microscopy (Figure 4M). We tracked their fate until the hatching stage (9 days postfertilization).

We found that the labeled cells contributed to blood vessels (presumably mural cells surrounding endothelial cells; Figure 4N, N’) and dorsal finfold mesenchyme (Figure 4O, O’), which is in accordance with previous studies revealing their somitic origins (Ando et al., Development 2016; Lee et al., Development 2013; Garriock and Krieg, 2007). However, we did not observe any labeled myotomal cells. These results suggest that these dorsal DM cells, guiding the myotome to cover the neural tube, do not contribute to the myotome themselves, at least at the hatching stage.

In the manuscript, we replaced the previous figures for fate mapping with these new figures (Figures 4M-O’) and described them in the results and Discussion sections as follows:

(Line 220, Results):

“Finally, we examined the fate of the dorsal DM cells that derive the mesenchymal cells. We employed a photoconversion technique mediated by a photoconvertible protein KikGR to specifically label a single dorsal DM cell exhibiting protrusions at the tip of a 10th or 20th somite during dorsal somite extension (Figure 4M, arrowhead) and tracked them until the hatching stage. We found that the labeled dorsal DM cells eventually differentiated into cells surrounding blood vessels (likely mural cells, n = 12/17 for 10th and 20th somites; Figure 4N, N’, arrowheads) and mesenchymal cells in the dorsal fin fold (n = 4/8 for 20th somites; Figure 4O, O’, arrowheads); however, we did not observe any labeled axial muscles (n = 0/17 for 10th and 20th somites). This suggests that the dorsal DM cells at the tip of somites, guiding myotome extension, do not become myotomal cells themselves at least until the end of embryonic development (hatching stage, 9 dpf).”

(Line 420, Discussion):

“Interestingly, our lineage tracing experiment revealed that the dorsal DM cells at the tip of somites differentiated into cells of non-myotomal lineages including mural cells of blood vessels and mesenchymal cells in the dorsal fin fold. These observations are consistent with previous studies reporting their somitic origins in zebrafish and Xenopus (Garriock and Krieg, 2007; Lee et al., 2013; Ando et al., 2016). However, our data also showed that these DM cells do not appear to contribute to the myotome at the hatching stage. We thus propose that the neural tube coverage is driven by a specific population of dorsal DM cells which do not contribute to myotomal cells during embryonic development.”

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

The manuscript has been improved but there are some remaining issues that need to be addressed, as outlined below:

The authors further improved the manuscript by adding fate mapping data of the dorsal DM cells. The fate mapping shows that the dorsally extending cells of interest contribute to the fin-fold mesenchyme and appear to surround blood vessels, however no labeled cells contribute to the myotome itself.

1) These new findings need to be better represented throughout the manuscript. For example, the abstract still reads:

'In wild type, dorsal dermomyotome (DM) cells, progenitors of myotomal cells, reduce their proliferative activity after somitogenesis and subsequently form unique large protrusions extending dorsally, guiding the epaxial myotome dorsally'

Please clarify that dorsal dermomyotome cells are not myotomal progenitors in this context.

We appreciate the reviewers’ suggestion and have modified the text to underline the difference between the commonly recognized role of DM cells as myotomal progenitors and our novel finding of the DM cells as a non-myogenic subpopulation as follows:

(Line 17, abstract): “In wild type, dorsal dermomyotome (DM) cells reduce their proliferative activity after somitogenesis. Subsequently, a subset of DM cells, which does not differentiate into the myotome population, begins to form unique large protrusions extending dorsally to guide the epaxial myotome dorsally.”

(Line 44, introduction): “While we have a detailed understanding of how the myotome, precursors of epaxial and hypaxial muscles, differentiates from a somitic compartment called the dermomyotome (DM)…”

(Line 83, introduction): “We also found that these DM cells form an subpopulation that gives rise to non-myotomal cell lineages during embryonic development.”

(Line 392, discussion): “DM cells have been known to serve as a progenitor pool for myotomal and dermal cells (Ben-Yair and Kalcheim, 2005; Hollway et al., 2007; Stellabotte and Devoto, 2007). In our study, we showed that DM cells at the tip of the dorsal somite form unique large protrusions, guiding the myotome towards the top of the neural tube. Furthermore, these DM cells do not give rise to myotomal cells during embryonic development.”

2) Given the new fate mapping data, please also make the modified model more explicit throughout the abstract, intro and discussion: that a non-myogenic lineage of pioneer cells non-cell-autonomously guides dorsal myotome growth.

Following the reviewers’ comment, we have highlighted the modified model involving the non-myogenic population of DM cells as shown below:

(Line 26, abstract): “We propose that dorsal extension of the epaxial myotome is guided by a non-myogenic subpopulation of DM cells and that wnt11 empowers the DM cells to drive the coverage of the neural tube by the epaxial myotome.”

(Line 89, introduction): “We thus propose an unprecedented process of epaxial myotome morphogenesis driven by a non-myogenic population of DM cells during embryogenesis.”

(Line 398, discussion): “We thus revealed a novel role of non-myogenic DM cells during epaxial myotome morphogenesis.”

3) Could the authors confirm that the fate mapping experiments consist of 17 individual cells in 17 different embryos?

In the fate mapping experiments, we mostly labeled individual DM cells in two different positions (10th and 20th somites) of single embryos and examined the effect of somite position on the myotomal contribution of DM cells. We labeled individual cells in 8 embryos at both 10th and 20th somites, and in 1 embryo we labeled an individual cell only at the 10th somite. During the experiment the labeled cells stayed in the same region during their differentiation, and we never observed movement of the labeled cells over a long distance to intermingle with other labeled cells from a different position.

To clarify this point, we modified the description on the sample size in the Results section (Lines 235, 238), as well as added a sentence in the Materials and methods (Line 556).

4) Title: 'Zic1 advances epaxial myotome morphogenesis to cover the neural tube via Wnt11'. Please consider rephrasing the title, which currently does not convey that wnt11 acts on DM cells, which then guide the epaxial myotome. The title suggests that wnt11 acts on epaxial myotome cells directly.

We have now changed the title to “Wnt11 acts on dermomyotome cells to guide epaxial myotome morphogenesis” to represent the role of Wnt11 more clearly.

We have also rewritten the impact statement to indicate the significance of our findings more explicitly.

“A medaka mutant revealed that Wnt11 promotes formation of uniquely large protrusions from non-myogenic dorsal dermomyotome cells, which guide the epaxial myotome dorsally to achieve the coverage of the neural tube.”

Line 571, 1371: CRISPR-mediated knock-out of wnt11. Please, rephrase knock-out as the analyzed embryos were not stable mutants. F0 CRISPR causes a hodgepodge of mutations and WT cells.

We fully agree with the reviewers’ comment and have rephrased the description as follows:

(Line 586, Materials and methods subtitle): “wnt11 sgRNA injection”

(Line 1373, Figure 7 —figure supplement 2 title): “Injection of wnt11 sgRNAs results in delayed epiboly movement, and impaired body axis and trunk development.”

Furthermore, we also added a few words to avoid confusion in the main text:

(Line 326): “Injection of single guide RNAs (sgRNAs) against wnt11 to knock out the gene revealed that the resultant genetically mosaic F0 embryos displayed delayed epiboly movement and subsequent morphological defects…”

Please soften the language regarding the morpholino experiments as initially requested by the reviewers.

Following the reviewers’ suggestion, we have now softened the expression of sentences claiming the function of wnt11 based on the morpholino experiments as below:

(Line 356): “Overall, knock-down of the Zic1 target gene wnt11 recapitulated the phenotype of Da DM cells (Figure 3G-G’’), suggesting the crucial role of Wnt11 in regulating protrusion formation of DM cells.”

(Line 451): “Our data suggest that in medaka dorsal somites Wnt11 exerts its effect through promotion of protrusion formation and down-regulation of cell proliferation in the dorsal DM.”

(Line 480): “Wnt11 may not be a sole factor for dorsal somite extension, although it was suggested to be crucial in the present study.”

https://doi.org/10.7554/eLife.71845.sa2

Article and author information

Author details

  1. Ann Kathrin Heilig

    1. Department of Biological Sciences, Graduate School of Science, University of Tokyo, Tokyo, Japan
    2. Centre for Organismal Studies, Heidelberg University, Heidelberg, Germany
    3. Heidelberg Biosciences International Graduate School, Heidelberg, Germany
    Contribution
    Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Validation, Visualization, Writing - original draft, Writing - review and editing
    Competing interests
    No competing interests declared
  2. Ryohei Nakamura

    Department of Biological Sciences, Graduate School of Science, University of Tokyo, Tokyo, Japan
    Contribution
    Data curation, Formal analysis, Funding acquisition, Investigation, Validation, Writing - review and editing
    Competing interests
    No competing interests declared
  3. Atsuko Shimada

    Department of Biological Sciences, Graduate School of Science, University of Tokyo, Tokyo, Japan
    Contribution
    Data curation, Investigation, Writing - review and editing
    Competing interests
    No competing interests declared
  4. Yuka Hashimoto

    Department of Biological Sciences, Graduate School of Science, University of Tokyo, Tokyo, Japan
    Contribution
    Data curation, Formal analysis, Investigation
    Competing interests
    No competing interests declared
  5. Yuta Nakamura

    Department of Biological Sciences, Graduate School of Science, University of Tokyo, Tokyo, Japan
    Contribution
    Resources
    Competing interests
    No competing interests declared
  6. Joachim Wittbrodt

    Centre for Organismal Studies, Heidelberg University, Heidelberg, Germany
    Contribution
    Supervision, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-8550-7377
  7. Hiroyuki Takeda

    Department of Biological Sciences, Graduate School of Science, University of Tokyo, Tokyo, Japan
    Contribution
    Conceptualization, Funding acquisition, Supervision, Writing - review and editing
    For correspondence
    htakeda@bs.s.u-tokyo.ac.jp
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-7932-6358
  8. Toru Kawanishi

    Department of Biological Sciences, Graduate School of Science, University of Tokyo, Tokyo, Japan
    Contribution
    Conceptualization, Funding acquisition, Investigation, Resources, Supervision, Writing - review and editing
    For correspondence
    toru.kawanishi@bs.s.u-tokyo.ac.jp
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7038-9769

Funding

Japan Society for the Promotion of Science (JP15H05859)

  • Hiroyuki Takeda

Japan Society for the Promotion of Science (JP19K23741)

  • Toru Kawanishi

Japan Society for the Promotion of Science (JP18K14620)

  • Ryohei Nakamura

Japan Science and Technology Agency (JPMJCR13W3)

  • Hiroyuki Takeda

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

Acknowledgements

We thank the members of the Takeda laboratory for constructive feedback and discussions on the project. We are greatful for Y Yamagichi and M Funato for fish husbandry. This work was supported by Japan Society for the Promotion of Science (JSPS) KAKENHI Grant Numbers JP15H05859 (HT), JP19K23741 (TK) and JP18K14620 (RN) and Japan Science and Technology Agency CREST Grant Number JPMJCR13W3 (HT).

Ethics

All experimental procedures and animal care were performed according to the animal ethics committee of the University of Tokyo (Approval No. 20-02).

Senior Editor

  1. Marianne E Bronner, California Institute of Technology, United States

Reviewing Editor

  1. Tatjana Piotrowski, Stowers Institute for Medical Research, United States

Publication history

  1. Received: July 1, 2021
  2. Preprint posted: July 12, 2021 (view preprint)
  3. Accepted: April 19, 2022
  4. Version of Record published: May 6, 2022 (version 1)

Copyright

© 2022, Heilig 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.

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  1. Ann Kathrin Heilig
  2. Ryohei Nakamura
  3. Atsuko Shimada
  4. Yuka Hashimoto
  5. Yuta Nakamura
  6. Joachim Wittbrodt
  7. Hiroyuki Takeda
  8. Toru Kawanishi
(2022)
Wnt11 acts on dermomyotome cells to guide epaxial myotome morphogenesis
eLife 11:e71845.
https://doi.org/10.7554/eLife.71845

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