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Dynein-Mediated Regional Cell Division Reorientation Shapes a Tailbud Embryo

Summary Regulation of cell division orientation controls the spatial distribution of cells during development and is essential for one-directional tissue transformation, such as elongation. However, little is known about whether it plays a role in other types of tissue morphogenesis. Using an ascidian Halocynthia roretzi, we found that differently oriented cell divisions in the epidermis of the future trunk (anterior) and tail (posterior) regions create an hourglass-like epithelial bending between the two regions to shape the tailbud embryo. Our results show that posterior epidermal cells are polarized with dynein protein anteriorly localized, undergo dynein-dependent spindle rotation, and divide along the anteroposterior axis. This cell division facilitates constriction around the embryo's circumference only in the posterior region and epithelial bending formation. Our findings, therefore, provide an important insight into the role of oriented cell division in tissue morphogenesis.


HIGHLIGHTS
Differently oriented cell division creates epithelial bending to shape the tail bud Epidermal cells divide in different orientations in two different adjacent regions Posterior cells undergo dynein-dependent spindle rotation and divide along A-P axis Dynein is enriched in the anterior surface of the posterior cells

INTRODUCTION
During animal development, tissues, organs, and organisms are shaped into specific structures. Elucidating the mechanisms of these dynamic processes at multiple levels-molecules, cells, tissues, and organisms-is an important and fundamental issue in developmental biology. Although the structures of animals show considerable variation in shape, recent research has demonstrated the involvement of some fundamental mechanisms in the process of creating tissues, organs, and organisms across animal species. An example of this is epithelial bending or invagination, which is found in several cell morphogenetic events including gastrulation in Drosophila, Xenopus, and sea urchin (Sawyer et al., 2010;Davidson, 2012), and formation of the tracheal system, dorsal transverse fold, and leg epithelial fold in Drosophila embryos (Kondo and Hayashi, 2013;Wang et al., 2012;Ambrosini et al., 2019). However, interestingly, studies have shown that epithelial bending is driven by different cellular mechanisms, including apical constriction (Sawyer et al., 2010;Martin and Goldstein, 2014), mitotic cell rounding (Kondo and Hayashi, 2013), differential positioning of adherence junctions (Wang et al., 2012), and an actomyosin structure connecting the apical surface to the nucleus, which is anchored to the basal side in apoptotic cells (Ambrosini et al., 2019).
Regulation of cell division orientation, a well-known mechanism for proper distribution of determinants during asymmetric cell divisions (Bergstralh et al., 2017;di Pietro et al., 2016), is also essential for tissue morphogenesis such as tissue elongation and thickening (Panousopoulou and Green, 2014;da Silva and Vincent, 2007;Lechler and Fuchs, 2005). This is achieved via controlling the organized spatial distribution of cells following the oriented cell divisions. Three main rules apply to regulation of cell division orientation. The first is the Hertwig's rule, wherein cells divide along the longest axis of the cell; this is frequently observed in epithelial cell divisions (Hertwig, 1884;Brun-Usan et al., 2017;Minc et al., 2011;Minc and Piel, 2012). The second is the Sachs' rule (Saches, 1878), typically found in early embryonic cell divisions during cleavage stages (Guerrier, 1970;Meshcheryakov and Beloussov, 1975), wherein cells divide in a direction perpendicular to the orientation of the previous division. The third is the cell polarization rule (Brun-Usan et al., 2017;Morin and Bellaïche, 2011), wherein cells are polarized either by external cues or localized intrinsic factors and divide according to the polarity. One of the molecular bases of such cell polarization is the core cortical machinery that orients mitotic spindles and is known to function in asymmetric cell divisions, including those of Drosophila neuroblasts (di Pietro et al., 2016) and the Caenorhabditis elegans P1 cell of the two-cell embryo (Grill and Hyman, 2005;Rose and Gö nczy, 2014). The core machinery comprises Mud, Pins, and Gai in Drosophila (Bowman et al., 2006;Schaefer et al., 2000); LIN-5, GPR1/2, and GOA-1/GPA-16 in C. elegans (Srinivasan et al., 2003;Colombo et al., 2003); and NuMA, LGN, and Gai in vertebrates (Du and Macara, 2004;Peyre et al., 2011). These proteins form a complex with the dynein motor at the cell cortex, thereby exerting a pulling force on the astral microtubules by moving along them until the spindle is properly aligned (di Pietro et al., 2016;Bergstralh et al., 2017).
Ascidians are marine invertebrates that belong to the phylum Chordata. They are useful model systems for studying cellular behavior during morphogenesis in embryonic stages because of their invariant cell lineage and small number of cells constituting the embryo (Conklin, 1905;Nishida, 1986;Kumano and Nishida, 2007;Ogura and Sasakura, 2013). One remarkable morphogenetic process during ascidian embryogenesis, in addition to neural tube closure (Hashimoto et al., 2015) and endoderm invagination during gastrulation (Sherrard et al., 2010), is tail formation. In transitioning from the neurula to tailbud stages, a boundary between the future trunk and tail regions appears morphologically as an hourglass-like epithelial bending at the middle level of the embryo along the anteroposterior (A-P) axis (Hotta et al., 2007, Figure 1A). Following this, the tail region undergoes extensive elongation along the A-P axis (Hotta et al., 2007, Figure 1A). This derives mainly from cell rearrangements (Munro andOdell, 2002a, 2002b;Passamaneck et al., 2007), rather than cell proliferation, and the tail region ends up being several times longer than is the trunk region. This is in contrast with other animals including some annelid and vertebrate species, in which tail formation depends on proliferation of a small group of cells known as posterior growth zones or tailbuds, which are located at the posterior end of the embryo (Williams and Nagy, 2017;McGrew et al., 2008).
In this study, we analyzed the molecular and cellular mechanisms that shape the tailbud embryo of the ascidian Halocynthia roretzi. We propose a model for epithelial bending and overall body shape mechanisms that require cell divisions oriented perpendicularly to each other in two different adjacent areas of the epithelial layer. We determined that epidermal cells in the anterior and future trunk region divided around the circumference of the embryo, whereas those in the posterior and future tail region divided along the A-P axis. We also found that the difference in division orientation between the anterior and posterior regions was established by dynein-mediated spindle rotation just before the cell divisions in the posterior cells, which resulted in its orientation along the A-P axis regardless of cell shape. Accordingly, this cell division results in constriction around the circumference of the embryo only in the posterior region and creates epithelial bending at the boundary of the anterior and posterior regions to shape the tailbud.

Cell Morphology around the Future Epithelial Bending Site
To understand the cellular mechanisms by which the hourglass-like epithelial bending to shape the H. roretzi tailbud embryo is generated on the lateral side of the neurula embryo ( Figure 1A), our first aim was to characterize the morphology of epidermal cells in the outermost and epithelial layer of the embryo during bending formation. Phallacidin staining of fixed neurula embryos revealed that epidermal cells around the future bending region exhibit rectangular shape, frequently arranged in two rows, with their long axes aligned around the circumference of the embryo (bracket in Figure 1B2). Cell lineage analyses, in which descendants of the anterior ectodermal cells named a4.2 were traced following injection of pairs of a4.2 cells with a fluorescent dye, Texas Red dextran, at the eight-cell stage ( Figure 1B1), revealed that the ectodermal A-P lineage boundary, which is defined by the second cleavage after fertilization, coincides Figure 1. Epithelial Bending Formation and Tailbud Shaping (A) Phallacidin-stained embryos at the neurula through early tailbud stages. The image of each stage is a single focal plane, where the area of the embryo is at maximum. A1-A4 are viewed with anterior to the left. Close-up views of the dot-lined square in A1-A4 are shown at the bottom (A1 0 -A4 0 ). (A1) At 1 h after the neurula rotation (NR+1 h) (Nishide et al., 2012), approximately 30 min before the start of the 11th epidermal cell divisions. (A2) At 1 h and 30 min after the neurula rotation (NR+1 h 30 min). (A3) 2 h after the neurula rotation (NR+2 h). (A4) 2 h and 45 min after the neurula rotation (NR+2 h 45 min). Arrowheads in A3 and A4 indicate the boundary between the trunk and tail regions. Scale bars, 100 mm (A1-A4), 20 mm (A1 0 -A4 0 ). (B1) Schematic diagram of eight-cell embryo injected for lineage trace. Lateral view, with anterior to the left. Fluorescent dye was microinjected into anterior ectodermal cells, named a4.2 cells, on both sides of the embryo. (B2) The descendants of a4.2 cells (magenta). Cell shape is visualized with Phallacidin (green). The boundary of anterior and posterior ectodermal lineages is indicated by an arrow. Rows of rectangular-shaped cells are indicated with white brackets. The digits in the bottom right-hand corner indicate the proportion of the embryo in which rows of rectangular-shaped cells were derived from a4.2 cells. Scale bar, 50 mm. (B3) Frontal section of an early tailbud embryo. The descendants of a4.2 cells (magenta), and cell shape is visualized with Phallacidin (green). The deepest position of the epithelial bending is indicated by an arrowhead. The digits in the bottom right-hand corner indicate the proportion of the embryo in which the lineage boundary corresponded to the deepest position of the epithelial bending. Scale bar, 50 mm. (C) Snap shot images obtained from live confocal imaging showing the epidermal cell division. The cell membrane was labeled with PH-YFP. Anterior is to the left. In each time point, a section along the anterior-posterior axis (top, X-Z section) and lateral view of the embryo (bottom, X-Y section) are shown. Shadow images of the X-Z section are shown for clarification of the formation of epithelial bending (red arrowheads). The dividing and divided cells in the X-Y plane are indicated with yellow and red dots, respectively. The digits in the bottom right-hand corner at the 10 min image indicate the proportion of the embryo in which epidermal cell division accompanied formation of epithelial bending. Scale bar, 100 mm.  Figure 1B2) and with the deepest point of the epithelial bending (89.5% [n = 19], arrowhead in Figure 1B3) following its formation.

Epidermal Cell Divisions during Epithelial Bending and Tailbud Shape Formation
We next carried out live-imaging analysis using PH-YFP encoding a fusion YFP fluorescent protein to a pleckstrin homology (PH) domain to visualize cell membranes (Tall et al., 2000;Passamaneck et al., 2007) of epidermal cells on the lateral side of the embryo. We found that the timing of the formation of epithelial bending (X-Z sections in Figure 1C) coincides with the timing of epidermal cell divisions (X-Y planes in Figure 1C). These cell divisions were the 11th to occur after fertilization ( Figure S1). To examine whether the epidermal cell divisions are related causally to the formation of epithelial bending, we cultured embryos in the presence of Hydroxyurea (HU), a cell division inhibitor, beginning at approximately 1 h and 30 min prior to the cell divisions. We found that HU treatment inhibited both cell division (bottom in Figure 1D) and formation of epithelial bending/tailbud shaping (top in Figure 1D) in a concentration-dependent manner. However, we cannot rule out the possibility that the resultant larger cell size itself might have blocked the morphogenesis. We also cannot eliminate the possibility that this treatment affects other cells, especially those located inside the embryo; however, previous studies suggest that at least notochord and muscle cells, which constitute a large portion of the tail region, have already ceased their embryonic cell divisions before the late neurula stage (Nishida, 1987;Kuwajima et al., 2014).

Differently Oriented Cell Divisions between Anterior and Posterior Regions
We then tried to clarify the potential contribution of those epidermal cell divisions to the formation of epithelial bending and the tailbud shaping. Via live-imaging analysis with PH-YFP, we found that epidermal cells undergo differently oriented cell divisions in anterior and posterior regions on the lateral side of the embryo. Cells in the anterior region divide around the circumference of the embryo (red lines in Figure 2A), whereas cells in the posterior region divide along the A-P axis (white lines in Figure 2A), forming a clear boundary of division orientation around the bending forming region (Figure 2A, Video S1). To ascertain the precise location of the division orientation boundary, we took advantage of asymmetric inheritance of the injected mRNA and identified the ectodermal A-P lineage boundary by means of different cell membrane fluorescence intensity. We ascertained that the division orientation boundary is located one cell row anterior to the A-P lineage boundary and that the one cell row difference was invariant in nearly all the embryos observed ( Figures 2B and 2C).
Comparison of division orientations between different rows of the rectangular-shaped cells (rows 1 and 2 in Figures 2A and 2B) is especially interesting as they divided in directions nearly perpendicular to each other (red circles in the middle two columns of Figure 2E), even though their cell geometries are almost identical in terms of cell shape and long axis orientation ( Figure 2D, black circles in the middle two columns of Figure 2E). The cells in row 1 divided along their long axes, whereas those in row 2 did not ( Figure 2E). Our analyses of other cells also revealed that epidermal cells in a row anterior to row 1 (row A in Figure 2B) divide (B) Schematic diagram of epidermal cells on the lateral side of the embryo. The cell row located anterior to the division orientation boundary is indicated as row 1, that posterior to row 1 is indicated as row 2, that anterior to row 1 is indicated as row A, that posterior to row 2 is indicated as row P, and that posterior to row P is indicated as row PP.
(C) Division orientation with respect to the A-P axis. The inset in the graph demonstrates how to measure the angle of division orientation with respect to the A-P axis. Each circle represents a measurement of a single cell. The values were extracted from 41 cells/8 embryos for row A, 58 cells/9 embryos for row 1, 62 cells/9 embryos for row 2, 41 cells/6 embryos for row P, and 23 cells/3 embryos for row PP. along their long axes (left-most column of Figure 2E), whereas epidermal cells in rows posterior to row 2 (rows P and PP in Figure 2B) do not (right-most column of Figure 2E). These findings suggest that cells in rows 2, P, and PP undergo cell divisions against the Hertwig's rule (Hertwig, 1884) and that a mechanism actively directs their division orientation so that they divide along the A-P axis.

Mitotic Spindle Rotation in Posterior Epidermal Cells
We next investigated how epidermal cells in the posterior region come to divide along the A-P axis. We conducted live-imaging analysis using MAP7-GFP, which is known to localize to the microtubules and spindle poles during mitosis (McDougall et al., 2015), together with PH-YFP, to visualize mitotic spindle orientation in epidermal cells. We found that the epidermal cells in rows 2, P, and PP undergo mitotic spindle rotation such that the spindles are oriented along the A-P axis before cell divisions (Figures 3A and 3B;Videos S2 and S3). In contrast, the spindles of the cells in row 1 remained oriented in the same direction and did not rotate ( Figure 3A; Video S2). Notably, the mitotic spindles were aligned along the long axes of the cells prior to rotation ( Figures 3A-3C). The spindles then rotated in either a clockwise or counterclockwise direction, with most of them achieving the reorientation via the shorter of the two rotation paths (76.3% [n = 38], Figure 3D).
Based on the above findings, we propose a model for the hourglass-like epithelial bending formation to shape the tailbud embryo of the ascidian H. roretzi ( Figure 3E). Epidermal cells on the lateral side of the neurula embryo divide in a different orientation perpendicular to each other in the anterior and posterior regions after the collective and regional rotation of mitotic spindles in the posterior cells. This cell division with different division orientation facilitates constriction around the embryo's circumference only in the posterior region, with a lesser number of daughter cells distributed around the circumference than in the anterior region, which leads to the generation of a bending at the boundary between the anterior and posterior regions.

Dynein-Mediated Mitotic Spindle Rotation and Tailbud Shape Formation
To evaluate our model, we tried to disrupt the mitotic spindle rotation pharmacologically and ascertain its effect on the formation of a thinner and longer posterior body part and epithelial bending for the tail bud shaping. Research has shown that a motor protein dynein localizes at a cell cortex and drives mitotic spindle rotation for determination of cell division orientation (Lu and Johnston, 2013;di Pietro et al., 2016). Therefore, we treated embryos with 30 mM of a dynein inhibitor, Ciliobrevin D (Firestone et al., 2012; Lecland and Lü ders, 2014), for 1 h before the 11th ectodermal cell divisions. This abrogated the division orientation of the epidermal cells in rows 2, P, and PP and directed it in a more circumferential direction, increasing the angles with respect to the A-P axis when compared with control cells treated with DMSO (Figures 4A-4C; Videos S4 and S5). The division angles of the cells treated with Ciliobrevin D, especially those in rows P and PP, were distributed relatively evenly in a wide range between 0 and 90 ( Figure 4C); this is likely because the angles of the cells' long axes in rows P and PP were distributed in a similar way ( Figure 2E), along which the mitotic spindles were aligned originally ( Figure 3C), and they divided according to Hertwig's rule. Intriguingly, the inhibitor-treated embryos did not have the normal shape of tailbud embryos and instead displayed a significantly reduced extent of epithelial bending, as evaluated using two iScience 23, 100964, March 27, 2020 different calculation methods ( Figure 4D). In addition, these embryos had wider and shorter tails than did normal embryos ( Figure 4E) (214.7 mm for the Ciliobrevin D-treated tail width versus 180.6 mm for control, p < 0.001, t test; 180.8 mm for the Ciliobrevin D-treated tail length versus 237.7 mm for control, p < 0.001, t test). In the normal epithelial bending formation, the tail becomes relatively thinner when compared with the trunk and longer ( Figure 4F). We also noticed that, in the Ciliobrevin D-treated embryos, cell divisions were delayed ( Figure S2). However, this delay is not likely a cause of the phenotype described above, because treatment with the inhibitor under different conditions (at 10 mM for 2 h) further delayed cell division but did not affect the shaping of the tailbud embryo ( Figure S2).
Along with Ciliobrevin D, we used another dynein inhibitor EHNA and observed its effect on the epithelial bending and tailbud shape formation. EHNA is reported to be less specific (Firestone et al., 2012), but it has been shown to disrupt mitotic spindle orientation in another ascidian species, Ciona intestinalis (Negishi and Yasuo, 2015). The EHNA treatment abrogated cell division orientation (Figures S3A and S3B; Video S6) in a manner similar to that with Ciliobrevin D treatment, with significantly less bending ( Figure S3C) and wider and shorter tail ( Figure S3D) when compared with control embryos; however, the morphologies of the resultant tailbud embryo seemed to differ between the two inhibitor treatments (Figures 4D and S3C). In fact, the EHNA-treated tail was significantly thinner and longer than the Ciliobrevin D-treated one (191.3 mm for the EHNA-treated tail width [ Figure S3D] versus 214.7 mm for the Ciliobrevin D-treated tail width [ Figure 4E], p < 0.001, t test; 215.0 mm for the EHNA-treated tail length [ Figure S3D] versus 180.8 mm for the Ciliobrevin D-treated tail length [ Figure 4E], p < 0.001, t test), possibly due to less inhibitory effect of EHNA on cell division orientation (a division angle of 42.2 on average with respect to the A-P axis for EHNA-treated cells of the row PP [ Figure S3B] versus 53.3 for Ciliobrevin D-treated PP cells [Figure 4C] and/or to the non-specific effect of EHNA).
These results suggest that the mitotic spindle rotation is dynein dependent and required for the tailbud shaping.

Anterior Enrichment of Dynein Protein in Posterior Epidermal Cells
We then investigated whether dynein protein is enriched in a localized cell cortical domain of posterior epidermal cells, as is shown in several asymmetrically dividing cells undergoing mitotic spindle rotations (Lu and Johnston, 2013;di Pietro et al., 2016). To observe sub-cellular localization of dynein protein, we first performed antibody staining using anti-dynein heavy chain antibody, which has been used successfully to locate dynein protein in early embryonic cells of another ascidian species, C. intestinalis (Negishi and Yasuo, 2015). The antibody staining suggested that dynein protein is present at the cell cortex and the peri-nuclear region of posterior epidermal cells ( Figure 5A). Significantly, signals for dynein at the cell cortex were more concentrated to the anterior edge of cells ( Figure 5A). The average ratio of the anterior to the posterior dynein signal intensities was 1.51 ( Figure 5B, n = 108, S.D. = 0.39). The anterior  localization was also confirmed by fluorescent observation using a fusion protein of dynein cytoplasmic intermediate chain (DCIC) to EGFP and PH-dTomato ( Figure 5C). Further confirmation of these findings was obtained by live-imaging analysis, wherein mRNAs encoding DCIC-EGFP and MAP7-mCherry were injected. EGFP signals accumulated at the cell cortex, presumably at the anterior cortex of the cell, given the earlier results, and in the peri-nuclear region ( Figure 5D; Video S7). Notably, the EGFP signals at the cell cortex were evident (cells P and PP with arrowheads in Figure 5D; Video S7) before the mitotic spindle rotation occurs (cells P and PP with arrowheads in Figure 5D; Video S7). These results suggest that posterior epidermal cells are polarized along the A-P axis with dynein protein accumulating at the cells' anterior edges and that the protein drives mitotic spindle rotation prior to cell division. We detected fluorescent signals suggesting anteriorly concentrated dynein protein accumulation in cells in row PP from both the antibody staining (60.0% [n = 30], Figure S4) and the live-imaging analysis (59.1% [n = 22], Figure 5D and Video S7); however, the percentages were small for cells in rows 2 (18.2% [n = 11]) and P (7.7% [n = 13]) from the antibody staining. In most of the cells in rows 2 and P, the protein was present at the cell cortex and showed no obvious bias in its localization along the A-P axis ( Figure S4).

Autonomous Cell Division Reorientation to the Epithelial Sheet
To obtain an insight into the mechanisms of cell division reorientation, we examined possible involvement of the underlying tissues such as muscle and notochord. We manually isolated the epidermal layer of the lateral side of the embryo from the underlying tissues approximately 40 min after neurula rotation (Nishide et al., 2012) (i.e., approximately 45 min before the cell divisions) and observed the patterns of division of epidermal cells in the isolated layer. We found that majority of the epidermal cells in rows 2, P, and PP in the isolated layer divided along the A-P axis, whereas the cells in rows A and 1 divided more in a circumferential direction (Figures 6A and 6B; Video S8), as observed in the intact embryos ( Figure 2). Therefore, underlying tissues do not appear to affect the division reorientation of epidermal cells in the posterior region, at least after the 40-min mark after neurula rotation (45 min prior to the cell divisions).
Notably, an initial step of convergent extension movement of the notochord occurred between 1 h 30 min and 2 h after neurula rotation ( Figures 6C and 6D), eliminating the possibility that cell division orientation in the overlying epidermal cells is affected by physical force generated by the notochord extension along the A-P axis.

DISCUSSION
In this study, we found a mechanism by which epithelial bending and overall tailbud body shape is generated during animal development. The mechanism for shaping an ascidian tailbud embryo centers on cell divisions, with division orientations between two adjacent regions of the epithelial layer directed perpendicular to each other. The differently oriented cell divisions in the adjacent regions produce a relative narrowing in one region and the layer ends up with a bending at their boundary.

Possible Other Mechanisms for the Epithelial Bending Formation
Although our present findings suggest that an ascidian tailbud embryo is shaped by the two-directional epithelial layer transformation by different cell division orientations, we cannot rule out the involvement of other mechanisms.
Initially, we suspected that an anisotropic apical constriction of epidermal cells at the future bending region may have a role in the formation of epithelial bending. The rectangular shape of the cells in that region (Figure 1B2) was reminiscent of the mesoderm precursor cells undergoing anisotropic apical constriction in the ventral midline of gastrulating Drosophila embryos (Dawes-Hoang et al., 2005;Fox and Peifer, 2007;Sawyer et al., 2010). However, for the following reasons, we concluded that this is not the case. First, in the frontal section of the embryos, the epidermal cells at the bending region did not display a wedge-like shape along the apico-basal axis ( Figures 1A3 and 1A4). Second, the deepest point of the epithelial bending did not coincide with the center point of the rectangular-shaped cell rows (in this case, the interface of rows 1 and 2) ( Figures 1B2 and 1B3). Coincidence with the center point would have been observed if bending is driven by anisotropic apical constriction.
Another mechanism that may contribute to the formation of epithelial bending is a shortening of the lateral sides of the epidermal cells along the apico-basal axis. A shortening of cells drives invagination of endoderm cells during gastrulation in another ascidian species (Sherrard et al., 2010). We observed that epidermal cells around the future bending site were extended along the apico-basal axis longer than were other epidermal cells located more anteriorly and posteriorly prior to the formation of bending (Figure 1A2) but their length was reduced to that of the other cells upon formation of the epithelial bending ( Figures 1A3 and 1A4). This could explain why there was still a small degree of bending in embryos treated with Ciliobrevin D ( Figure 4D). Moreover, EHNA treatment reduced epithelial bending by a significantly greater extent than did Ciliobrevin D according to one of our calculation methods (8.18 mm in Figure 4D [Ciliobrevin D] versus 6.34 mm in Figure S3C [EHNA] calculated by Method 2, p < 0.001 [t test]) despite the fact that it still generated longer and thinner tails than did Ciliobrevin D (Figures 4 and S3). EHNA has been reported to be less specific to dynein inhibition and to interfere with actin-dependent cellular processes (Schliwa et al., 1984). Consequently, the EHNA treatment may have affected the shortening of the epidermal cells through inhibiting actin function and reducing the extent of the epithelial bending.
It may also be possible that tissues underlying the epidermal layer, such as muscle and notochord, are involved in the formation of epithelial bending and the tailbud shaping. Although our isolation experiment of the epidermal layer suggests that epidermal cells undergo division reorientation without the underlying tissue ( Figure 6A), it is still possible that underlying tissues are involved in ways other than regulation of cell division orientation. Suggestively, the boundary between underlying tissues, namely, mesenchyme and muscle, appears to coincide with the epithelial bending position.

Environmental Factors Required for the Tailbud Shaping
Our model for shaping the ascidian tailbud embryo ( Figure 3E) would be valid if the field around the embryo's circumference is resistant to a tension generated in the posterior region by the differently oriented cell divisions in the two adjacent regions ( Figure S5A, top). Accordingly, the constriction in the posterior region would properly take place as a result of the cell divisions. We speculate that the neural tube closure (Hashimoto et al., 2015; Ogura and Sasakura, 2016) could provide such a field, as we had found that the neural tube was nearly closed (two-thirds to three-quarters from the posterior end) when the epidermal cells divided (100% [n = 46]). If the neural tube is not closed, it is likely that the tension generated by the differently oriented cell division would not be propagated normally around the circumference of the embryo and posterior region would not be constricted ( Figure S5A, bottom). Although our current approaches (see the Limitations of the Study section) are inconclusive, it is tempting to propose the importance of ''a chain of morphogenetic events'' for the correct shaping of individual tissues in the scale of the whole organism, in which one event (in this case, neural tube closure) influences another event (tailbud shaping) occurring at a different place and time. Recently, it has been reported that neural tube closure is necessary for correct orientation of cell division and epidermal stratification in mouse embryos (Box et al., 2019). Consequently, future studies will be required to examine whether the interactive regulation of spatially and temporally different morphogenetic events is a common characteristic in animal morphogenesis.

Determination of the Position of the Division Orientation Boundary
Our present findings suggest that determination of the position of the epithelial bending along the A-P axis in the neurula embryo relies on the position of the cell division orientation boundary. We speculate that the position of the division orientation boundary along the A-P axis is determined by the cell lineage, rather than by extracellular signals occurring at a later stage-such as antagonistic signals for patterning the neurula embryo along the A-P axis (Pasini et al., 2012)-because the division orientation boundary fell consistently at the same position with respect to the A-P lineage boundary in the ectoderm (i.e., one cell row away anteriorly) in different individuals ( Figures 2B and 2C). As early as the second cell cleavage after fertilization that separates the A-P lineage, a difference could be made between the two lineages and memorized until it comes into effect at the late neurula stage. One candidate for the identity of the difference could be a member(s) of Postplasmic/Pem RNAs, a group of maternal mRNAs that are localized to the future posterior region of the egg and are inherited by posterior cells during cleavage stages (Makabe and Nishida, 2012;Sardet et al., 2005;Prodon et al., 2007). Although previous studies on Postplasmic/ Pem RNAs have identified their function in determination of developmental fate (Nishida and Sawada, 2001;Nakamura et al., 2006) and regulation of cell division patterning (Negishi et al., 2007;Prodon et al., 2010) in the posterior-vegetal cells (mesoderm and endoderm), one member named Pem has been shown to repress anterior ectodermal genes, such as FoxA and SoxB1, in posterior-animal cells (ectoderm) at the eight-cell stage (Kumano et al., 2011).
Currently, it is also unclear as to how the division orientation boundary is placed one cell row anterior to the A-P lineage boundary. We determined that posterior epidermal cells are polarized along the A-P axis, and, therefore, planar cell polarity (PCP) pathway may contribute to establishing cell polarization. PCP is known to be propagated in the Drosophila wing disc, from the prospective proximal region to the prospective distal region, to pattern the orientation of wing hair (Ambegaonkar et al., 2012;Devenport, 2014). In the ascidian Ciona savignyi, a polarizing signal has been shown to be propagated from a notochord cell to establish PCP in an adjacent notochord cell located anteriorly (Kourakis et al., 2014). Based on these findings, we speculate that, in the H. roretzi neurula embryo, the epidermal cells in row 2, the only cells that undergo spindle rotations derived from the anterior ectoderm lineage, may receive a polarizing signal from the cells derived from the posterior ectoderm lineage. However, we observed that the majority of the epidermal cells in row PP are polarized in terms of localization of dynein protein (Figures 5 and S4; Video S7), whereas those in rows 2 and P are not ( Figure S4). Therefore, the cells that share the boundary of the A-P lineages, both those in row 2 that share it posteriorly and those in row P that share it anteriorly, may behave differently owing to their contact with the different lineage cells across the boundary. However, they still undergo rotation of the mitotic spindle in a dynein-dependent manner ( Figures 4A, 4B, S3A, and S3B; Videos S5 and S6).

Shaping of the Tailbud Embryo in Another Ascidian Species
The shape of the tailbud embryo is conserved in different ascidian species, with epithelial bending formed in the middle of the embryo along the A-P axis and the tail region becoming thinner and longer (Hotta et al., 2007). However, study has shown that most of the epidermal cells in both the anterior and posterior regions of the neurula embryo in the ascidian C. intestinalis type A (Currently C. robusta) divide along the A-P axis at their 11th divisions (Ogura et al., 2011;Negishi et al., 2016). In addition, live-imaging observation indicated that the timing of an epithelial bending to shape the tailbud embryo of Ciona seemed not to coincide with the 11th cell divisions (Negishi et al., 2016;Negishi, personal communication). Therefore, H. roretzi and C. intestinalis type A (Currently C. robusta) may use different mechanisms for shaping the tailbud embryo and may exemplify a phenomenon known as developmental system drift, whereby divergence of underlying molecular mechanisms occurs for conserved tissue development across species (Lemaire and Piette, 2015;True and Haag, 2001).

Limitations of the Study
As discussed above, the relationship between neural tube closure and tailbud shaping is still speculative. At present we have tried to address this issue by the following approaches.
We prevented the neural tube from closing by removing the chorion, as has been reported (Nishida and Satoh, 1985), at the late gastrula stage, and determined, via the four calculation methods described earlier, that such embryos failed to shape the normal tailbud embryos with a reduced extent of epithelial bending and wider and shorter tail (Figures S5B and S5C). However, these embryos exhibited the normal pattern of cell division-around the circumference of the embryo in the anterior versus along the A-P axis in the posterior regions (Figures S5D and S5E; Video S9)-suggesting that the failure is not due to an abrogated pattern of cell division.
We also tried to make the field around the embryo's circumference less resistant to a tension generated by cell division by culturing embryos in Ca 2+ -free sea water, thereby weakening cell-cell adhesion. From right after the neural tube had been closed to two-thirds but before cell division, embryos were cultured in the absence of Ca 2+ . These embryos exhibited a reduced extent of epithelial bending and wider and shorter tail ( Figures S6A and S6B) but normal cell division orientations (Figures S6C and S6D).
These results support our speculation that a resistant field to the tension around the circumference of the embryo created by the neural tube closure could be necessary for shaping the epithelial bending and the tailbud embryo. However, non-specific effects cannot be ruled out in these experiments.
In another ascidian C. intestinalis type A (currently C. robusta), neural tube closure was successfully blocked by targeted overexpression of a dominant-negative form of RhoA (Hashimoto et al., 2015) and Cadherin 2 (Hashimoto and Munro, 2019) to dorsal midline cells including neural and epidermal cells flanking the boundary of the both cell types. Therefore, for the future prospective, it would be worthwhile to evaluate the epithelial bending and tailbud shape formation in H. roretzi embryos upon such more specific treatments to disrupt neural tube closure. These studies would shed light on an interactive regulation of spatially and temporally different morphogenetic events and its importance for tissue morphogenesis.

Supplemental Information
Dynein-Mediated Regional Cell Division

TRANSPALENT METHODS Animals and embryos
We obtained adults of Halocynthia roretzi from local fishermen near the Asamushi Research Center for Marine Biology (Aomori, Japan). Naturally-spawned eggs were fertilized using a suspension of nonself sperm. Embryos were cultured in Millipore-filtered seawater (MFSW) containing 50 µg/ml streptomycin and 50 µg/ml kanamycin at 9-13°C.

Constructs
This study used the following constructs: DCIC-EGFP: mRNA synthesis and microinjection mRNAs encoding the fusion proteins (DCIC-EGFP, MAP7-GFP, MAP7-mCherry, H2B-GFP, PH-YFP, PH-dTomato) were synthesized in vitro using the mMessage mMachine kit (Ambion), and poly(A) was added to the 3' ends of the synthesized mRNAs using the Poly(A) Tailing kit (Ambion) according to the manufacturer's instructions. We used the following restriction enzymes and RNA polymerases: T3 and NotI for DCIC-EGFP; T3 and SfiI for MAP7-GFP; T3 and NotI for MAP7-mCherry; T3 and PstI for H2B-GFP; T3 and NotI for PH-YFP; and T3 and SfiI for PH-dTomato. The synthesized mRNA was microinjected into fertilized eggs, a4.2, b4.2, a7.15, or b7.15 cells according to Miya et al. (1997) and Niwano et al. (2009) as described in the following. Fertilized eggs were treated with 0.05% actinase E (Kaken Pharmaceutical, Japan) in MFSW for 10 min to remove the follicle cells on the chorion. The treated eggs were washed with MFSW and transferred on the cover glass in plastic petri dish filled with MFSW. The eggs with chorion but without follicle cells were stuck on the cover glass. The solution to be microinjected was loaded in glass needles for microinjection which were made by pulling glass capillaries (GDC-1, Narishige) with a horizontal puller (PN-3, Narishige). Microinjection was carried out with a micromanipulator (MN-151, Narishige) under the dissection microscope (OLYMPUS SZX16).

Live imaging
Time lapse observation was performed using either a LSM710 Zen zeiss confocal microscope (Fig. 1C, 3A, 3B, Supplemental Movies 2 and 3) or a Nikon eclipse Ti ( Fig. 2A, 4A, 5D, 6A, Supplemental Movies 1, 4-9). For observation with the LSM710 Zen zeiss confocal microscope, embryos with vitelline membrane were mounted on the slide glass in random orientations, and well-oriented embryos were selected for observation of their lateral sides. Images were obtained every 2 min at 1.0-2.5 µm intervals. For observation with the Nikon eclipse Ti, the vitelline membranes were removed with forceps in a plastic dish coated with 1% agarose in MFSW. The embryos were transferred to a glassbottomed dish (Matsunami) filled with 2% methylcellulose in MFSW and carefully oriented using tungsten needles so that the lateral side could be observed. Images were obtained every 2-4 min at 4.0 µm intervals and were then subjected to deconvolution with the Nikon NIS-Elements software to reduce blurriness. The two different methods revealed the same pattern of epidermal cell divisions.

Drug treatment
Dynein inhibitor Ciliobrevin D (Calbiochem) was dissolved in DMSO at 50 mM and stored at −20°C as a stock solution. Before use, the stock solution was diluted to 30 µM in MFSW. The embryos were treated with 30 µM Ciliobrevin D for 1 h, beginning 1 h after neurula rotation (NR) (approximately 30 min before the 11th epidermal cell divisions). In another experiment, embryos were treated with 10 µM Ciliobrevin D for 2 h, beginning 1 h after NR. Following the treatment, embryos were washed with MFSW and then either used for live-imaging, as described above, or placed under the dissection microscope (OLYMPUS SZX16) to observe the epithelial bending. The 11th cell divisions had not begun at the time of washing with MFSW. The control embryos were treated with 0.4% DMSO for 1h from the NR+1 h stage. Another dynein inhibitor, EHNA (Santa Cruz Biotechnology), was dissolved in sterilized water at 50 mM and stored at 4°C as a stock solution. Before use, the stock solution was diluted in MFSW to either 1 mM or 2 mM. At 1 h after NR, the embryos were treated with 2 mM EHNA for 1 h and then treated with 1 mM EHNA for another 30 min. The treated embryos were washed with MFSW and used for live-imaging, as described above. Hydroxyurea (HU) (Sigma) was dissolved in MFSW at 20 mM, 50 mM and 100 mM.

Definition of the start and the end of the 11th cell division
When epidermal cells begin to divide, the surface of the embryo becomes rough due to the formation of division furrows. This can be detected by the dissection microscope (OLYMPUS SZX16) and is defined as the onset of the 11th cell division. The surface of the embryo becomes smooth upon completion of epidermal cell divisions. This can also be detected by the dissection microscope and is defined as the end of the 11th cell division.

Immunohistochemistry and Phallacidin staining
For anti-dynein antibody staining, embryos were fixed in ice-cold methanol for 10 min and washed four times with 0.1% Triton X-100 in PBS (PBSTr). The fixed embryos were treated with 3% H 2 O 2 in PBS for 10 min and with 50 mM NH 4 Cl in PBS for 15 min. After being washed with PBS four times, the specimens were incubated for 30 min at room temperature with 0.5% blocking reagent (Roche) in PBS. Anti-dynein heavy chain antibody (R-325, Santa Cruz Biotechnology) was diluted at 1:50 in 0.5% blocking reagents in PBS, and the specimens were incubated overnight at 4°C. The primary antibody was washed with PBSTr four times. We used the MaxPO system (Nichirei Corporation) as a secondary antibody. Anti-rabbit antibody conjugated with HRP [MAX-PO (R)] was diluted at 1:20 in 0.5% blocking reagents in PBS, and the specimens were incubated overnight at 4°C. The secondary antibody was washed four times with PBSTr, and HRP activity was detected using the TSA Plus Cy3 kit (Perkin-Elmer Life Sciences) according to the manufacturer's instruction.
To identify the lineage boundary, H2B-GFP mRNA was injected into the b4.2 cell at the 8-cell stage, and the embryos were fixed as described above. The fixed embryos were double stained with antidynein heavy chain antibody and anti-GFP antibody (ab13970, Abcam), as described above, with the following exception: anti-GFP antibody was diluted at 1:200 in 0.5% blocking reagents in PBS. As a secondary antibody, anti-chicken Alexa 488 antibody (A11039, Invitrogen) was diluted at 1:100 in 0.5% blocking reagents in PBS.
To determine which side of the cell was enriched with dynein protein, specimens were squished gently to separate the cells, and cell boundaries were identified using the differential interference contrast (DIC) optics of the Zeiss LSM5 PASCAL confocal microscope. For Phallacidin (Thermo Fisher Scientific) staining, embryos were fixed for 30 min in 4% paraformaldehyde in a buffer containing 50 mM EGTA, 100 mM Pipes and 400 mM sucrose, adjusted to pH 6.9 (Munro and Odell, 2002a). The fixed embryos were washed three times with PBS and incubated with Phallicidin (5 unit/ml) in PBSTr either for 1 h at room temperature or overnight at 4°C and then washed three times with PBS. The embryos were treated with isopropanol series (70%, 85%, 95% and 100% in PBS) for 1 min each, and cleared with Murray clear (Benzyl Benzoate:Benzyl Alcohol = 2:1). The specimens were observed using the Zeiss LSM5 PASCAL confocal microscope.

Isolation of epidermal cell sheet
The vitelline membrane was removed manually using forceps, and the embryos were transferred to plastic dish coated with 1% agarose in MFSW. Embryos were dissected along the midline using an eyebrow hair mounted on the tip of a glass needle. Either the left side half or the right side half was used. The tissues underlying muscle and notochord cells were identified by their large cell size and cell shape. For removal of the dorsal and ventral regions, the locations of the muscle and notochord cells were employed as landmarks of the lateral region. After the removal of dorsal and ventral epidermal cells, the underlying muscle and notochord cells were removed gently with the eyebrow hair. The isolated ectodermal cell sheets were transferred gently to 2% methylcellulose in MFSW and the pattern of division was observed using the Nikon eclipse Ti, as described above.