Author response:
The following is the authors’ response to the original reviews.
Public Reviews:
Reviewer #1 (Public review):
Summary:
This paper investigates the physical basis of epithelial invagination in the morphogenesis of the ascidian siphon tube. The authors observe changes in actin and myosin distribution during siphon tube morphogenesis using fixed specimens and immunohistochemistry. They discover that there is a biphasic change in the actomyosin localization that correlates with changes in cell shapes. Initially, there is the well-known relocation of actomyosin from the lateral sides to the apical surface of cells that will invaginate, accompanied by a concomitant lengthening of the central cells within the invagination, but not a lot of invagination. Coincident with a second, more rapid, phase of invagination, the authors see a relocalization of actomyosin back to the lateral sides of the cells. This 2nd "bidirectional" relocation of actin appears to be important because optogenetic inhibition of myosin in the lateral domain after the initial invaginations phase resulted in a block of further invagination. Although not noted in the paper, that the second phase of siphon invagination is dependent on actomyosin is interesting and important because it has been shown that during Drosophila mesoderm invagination that a second "folding" phase of invagination is independent of actomyosin contraction (Guo et al. elife 2022), so there appear to be important differences between the Drosophila mesoderm system and the ascidian siphon tube systems.
Using the experimental data, the authors create a vertex model of the invagination, and simulations reveal a coupled mechanism of apicobasal tension imbalance and lateral contraction that creates the invagination. The resultant model appears to recapitulate many aspects of the observed cell behaviors, although there are some caveats to consider (described below).
We thank the reviewer for the insightful summary and for bringing the important study by Guo et al. (2022) to our attention. We have now added a dedicated comparison with Drosophila ventral furrow invagination in the Discussion, explicitly highlighting that the second rapid folding phase in Drosophila does not require lateral contractility, whereas in our system lateral contractility is obligatory for the accelerated invagination stage.
Strengths:
The studies and presented results are well done and provide important insights into the physical forces of epithelial invagination, which is important because invaginations are how a large fraction of organs in multicellular organisms are formed.
Thank you for this positive assessment and for recognizing the significance of our work in elucidating the physical mechanisms underlying fundamental morphogenetic processes. We have striven to provide a comprehensive and rigorous analysis, and are grateful for this encouraging feedback.
Weaknesses:
(1) This reviewer has concerns about two aspects of the computational model. First, the model in Figure 5D shows a simulation of a flat epithelial sheet creating an invagination. However, the actual invagination is occurring in a small embryo that has significant curvature, such that nine or so cells occupy a 90-degree arc of the 360-degree circle that defines the embryo's cross-section (e.g., see Figure 1A). This curvature could have important effects on cell behavior.
Thank you for bringing up the issue of tissue curvature. In the initial version of our model, we treated the tissue as flat based on the local geometry of the anterior epidermis. Although the embryo at 13 hpf indeed possesses significant curvature, its overall transverse cross-section is approximately elliptical, and the region undergoing invagination is situated in a relatively low-curvature zone, occupying only a 30° ∼ 40° arc of the entire tissue. More importantly, the embryo undergoes anisotropic elongation and expansion, becoming significantly flattened during the accelerated invagination stage, eventually adopting a very flat geometry by 18 hpf. We have now included Figure 5—figure supplement 1 to clarify these global morphological transitions.
Nevertheless, the curvature does exist during the early stages, and we agree that clarifying its potential role is essential. Therefore, in the revised manuscript, we have updated our vertex model to incorporate a simplified circular geometry. Furthermore, unlike Drosophila ventral furrow formation (Guo et al., eLife, 2022), the invagination here eventually forms a hollow tubular structure, which led us to introduce a surface bending stiffness term into the mode. Although global tissue growth is not explicitly modeled, we explored the impact of curvature by varying the initial system size. Our results demonstrate that the invagination process, driven by apico-basal tension imbalance and lateral contraction, is highly localized and remains robust across different curvatures.
(2) The second concern about the model is that Figure 5 D shows the vertex model developing significant "puckering" (bulging) surrounding the invagination. Such "puckering" is not seen in the in vivo invagination (Figure 1A, 2A). This issue is not discussed in the text, so it is unclear how big an issue this is for the developed model, but the model does not recapitulate all aspects of the siphon invagination system.
Thank you for pointing out this. In our experiments, the similar "puckering" shape is observed during the early stages of morphogenesis (~17 hpf, as seen in Figure 1A) when the tissue size is relatively small. However, this feature rapidly disappears as the tissue grows and the overall geometry becomes flatter. This suggests that "puckering" is more pronounced in highly curved epithelia, a phenomenon that aligns with mechanical expectations. Previous vertex models of Drosophila ventral furrow formation do not exhibit this effect (Brodland et al., 2010; Polyakov et al., 2014), because they modeled cells within a rigid unmovable boundary. However, in our system of siphon morphogenesis, a tubular structure ultimately forms in the epithelium without strong boundary constraints. Thus, the mechanical boundary conditions are basically different.
Also, the formation of a hollow tubular structure—supported by strong F-actin accumulation at the tissue surface—indicates a bending stiffness of surface tissue (Figure 1), which we have incorporated into the model. This bending term enforces smooth curvature transitions, which can manifest as a "puckering" shape surrounding the invagination. In our previous flat-geometry model, this significant bending stiffness led to a "puckering" effect surrounding the invagination. In our updated curved vertex model, this phenomenon also exists and is found to be related to tissue curvature. By simulating a larger system with low curvature (N = 324 cells in Figure 6D), we find that this puckering is significantly reduced. This confirms that the shape discrepancy is a size-dependent effect of the bending constraints within a fixed system size that did not account for tissue growth. In biological development, continuous growth and flattening of the embryo diminish this effect (Figure 5—figure supplement 1), aligning our model's predictions.
Furthermore, we note that the cell-cell adhesion between the surface epithelium and the internal bulk cells (a factor not explicitly captured in our current model) likely further suppresses such evagination in vivo, as outward puckering would necessitate the coordinated deformation of the underlying tissues. We aim to investigate the interplay between global growth and local active forces in future work. We have added a detailed description and mechanical explanation of these simulated shapes in the revised manuscript.
(3) In Figure 2A, Top View, and the schematic in Figure 2C, the developing invagination is surrounded by a ring of aligned cell edges characteristic of a "purse string" type actomyosin cable that would create pressure on the invaginating cells, which has been documented in multiple systems. Notably, the schematic in Figure 2C shows myosin II localizing to aligned "purse string" edges, suggesting the purse string is actively compressing the more central cells. If the purse string consistently appears during siphon invagination, a complete understanding of siphon invagination will require understanding the contributions of the purse string to the invagination process.
Thank you for this excellent observation. We agree that the ring-like actomyosin structure is a prominent feature during the initial stages of invagination, and its potential role warrants discussion. We carefully re-examined our data. Our analysis confirms that this myosin ring is most pronounced during the early initial invagination stage. This inward compression from the periphery would work in concert with apical constriction to help shape the initial invagination. However, this ring-like myosin pattern significantly diminishes during the accelerated invagination stage, indicating that sustained compression from the purse string is not required for the entire process. We have added a discussion of this point in the revised manuscript. We also agree with that future experiments using laser ablation or optogenetic inhibition specifically targeting this actomyosin ring would be valuable to further dissect its precise contribution during the early invagination stage, and we have noted this as a future direction in the Discussion.
(4) The introduction and discussion put the work in the context of work on physical forces in invagination, but there is not much discussion of how the modeling fits into the literature.
We thank the reviewer for this suggestion. We have now incorporated additional references and discussion regarding existing theoretical models and the physical forces involved in tissue invagination. These previous studies provided the foundational framework for our updated curved vertex model. We have also added an explanation of how our model differs from these existing works and discussed potential future directions for further investigation.
Reviewer #2 (Public review):
Summary:
The authors propose that bidirectional translocation of actomyosin drives tissue invagination in Ciona siphon tube formation. They suggest a two-stage model where actomyosin first accumulates apically to drive a slow initial invagination, followed by translocation to lateral domains to accelerate the invagination process through cell shortening. They have shown that actomyosin activity is important for invagination - modulation of myosin activity through expression of myosin mutants altered the timing and speed of invagination; furthermore, optogenetic inhibition of myosin during the transition of the slow and fast stages disrupted invagination. The authors further developed a vertex model to validate the relationship between contractile force distribution and epithelial invagination.
Thank you for your thoughtful and accurate summary of our work and for your constructive critique.
Strengths:
(1) The authors employed various techniques to address the research question, including optogenetics, the use of MRLC mutants, and vertex modelling.
(2) The authors provide quantitative analyses for a substantial portion of their imaging data, including cell and tissue geometry parameters as well as actin and myosin distributions. The sample sizes used in these analyses appear appropriate.
(3) The authors combined experimental measurements with computer modeling to test the proposed mechanical models, which represents a strength of the study. It provides a framework to explore the mechanical principles underlying the observed morphogenesis.
We are grateful for your positive assessment of the multidisciplinary approaches, quantitative analyses, and the integration of modeling with experiments.
Weaknesses:
(1) The concept of coordinated and sequential action of apical and lateral actomyosin in support of epithelial folding has been documented through a combination of experimental and modeling approaches in other contexts, such as ascidian endoderm invagination (PMID: 20691592) and gastrulation in Drosophila (PMIDs: 21127270, 22511944, 31273212). While the manuscript addresses an important question, related findings have been reported in these previous studies. This overlap reduces the degree of novelty, and it remains to be clarified how their work advances beyond these prior contributions.
We thank the reviewer for raising this important point. In the revised Introduction and Discussion, we have explicitly distinguished our findings from prior studies. Specifically: (1) Unlike ascidian endoderm invagination, where actomyosin shifts from apical to basolateral (Sherrard et al., 2010), our system exhibits a bidirectional redistribution between apical and lateral domains, with the basal domain playing a passive role. (2) Unlike Drosophila ventral furrow invagination, where lateral contractility is not essential for the second folding phase (Guo et al., 2022), our optogenetic inhibition demonstrates that lateral contractility is obligatory for the accelerated invagination stage. These comparisons, now clearly stated in the Introduction and Discussion, establish bidirectional actomyosin redistribution as a distinct mechanical paradigm for sequential morphogenesis. We believe these revisions adequately clarify how our work advances beyond prior contributions.
(2) One of the central statements made by the authors is that the translocation of actomyosin between the apical and lateral domains mediates invagination. The use of the term "translocation" infers that the same actomyosin structures physically move from one location to another location, which is not demonstrated by the data. Given the time scale of the process (several hours), it is also possible that the observed spatiotemporal patterns of actomyosin intensity result from sequential activation/assembly and inactivation/disassembly at specific locations on the cell cortex, rather than from the physical translocation of actomyosin structures over time.
We thank the reviewer for this important point. We agree that our data do not demonstrate physical translocation of actomyosin structures, and that the observed patterns could arise from sequential assembly/disassembly over time. To avoid overinterpretation, we have replaced “translocation” with “redistribution” throughout the manuscript (including the title) and toned down the language in the Results and Discussion.
(3) Some aspects of the data on actomyosin localization require further clarification. (1) The authors state that actomyosin translocation is bidirectional, first moving from the lateral domain to the apical domain; however, the reduction of the lateral actomyosin at this step was not rigorously tested. (2) During the slow invagination stage, it is unclear whether myosin consistently localizes to the apical cell-cell borders or instead relocalizes to the medioapical domain, as suggested by the schematic illustration presented in Figure 2C. (3) It is unclear how many cells along the axis orthogonal to the furrow accumulate apical and lateral myosin.
Thank you for your insightful comments, which will help us significantly improve the clarity and rigor of our actomyosin localization analysis. To address the points raised, we undertake several key revisions: First, we have added new quantitative analyses of active myosin intensity from earlier time points (14-15 hpf) to rigorously support the initial lateral-to-apical redistribution phase (Figure 2B). Second, the schematic in Figure 2C has been corrected to show myosin at the apical cell‑cell borders. We have clarified that redistribution occurs in a domain of approximately 15‑20 cells (the invagination primordium), not only the center cell.
(4) The overexpression of MRLC mutants appears to be rather patchy in some cases (e.g., in Figure 3A, 17.0 hpf, only cells located at the right side of the furrow appeared to express MRLC T18ES19E). It is unclear how such patchy expression would impact the phenotype.
Thank you for your observation. We acknowledge that mosaic expression is common in Ciona electroporation. For all quantitative analyses, we only selected embryos in which the central cell, along with more than half of the surrounding cells in the primordium, showed clear expression of the plasmid. This selection criterion has been added to the Materials and Methods section.
(5) In the optogenetic experiment, it appears that after one hour of light stimulation, the apical side of the tissue underwent relaxation (comparing 17 hpf and 16 hpf in Figure 4B). It is therefore unclear whether the observed defect in invagination is due to apical relaxation or lack of lateral contractility, or both. Therefore, the phenotype is not sufficient to support the authors' statement that "redistribution of myosin contractility from the apical to lateral regions is essential for the development of invagination".
We have performed the additional immunostaining experiment of myosin II. The new data (Figure 4—figure supplement 2) showed that light stimulation specifically reduced lateral myosin intensity without significantly affecting apical myosin compared to the dark control. Therefore, the observed block of invagination is primarily due to loss of lateral contractility.
(6) The vertex model is designed to explore how apical and lateral tensions contribute to distinct morphological outcomes. While the authors raise several interesting predictions, these are not further tested, making it unclear to what extent the model provides new insights that can be validated experimentally. In addition, modeling the epithelium as a flat sheet and not accounting for cell curvature is a simplification that may limit the model's accuracy. Finally, the model does not fully recapitulate the deeply invaginated furrow configuration as observed in a real embryo (comparing 18 hpf in Figure 5D and 18 hpf in Figure 1A) and does not fully capture certain mutant phenotypes (comparing 18 hpf in Figure 5F and 18 hpf in Figure 3B right panel).
Thank you very much for these helpful and constructive comments. We have addressed your concerns through the following model updates and clarifications.
First, we have reformulated our vertex model from a flat sheet to a curved geometry that incorporates initial tissue curvature. We found that the core mechanical mechanism, mediated by the coupling of apical and lateral active contraction, consistently recapitulates the experimental invagination process. By independently inhibiting apical or lateral contractions in the model, we further clarified their distinct mechanical contributions to tissue bending and cell shortening.
Regarding the model predictions concerning the apical-to-lateral redistribution of actomyosin in the original version (previously shown in Figure 6E-H), we agree that these lacked direct experimental validation in the current study and may have strayed from the primary focus on the invagination mechanism itself. Therefore, we have removed these predictive components from the revised manuscript. Instead, we have refocused our analysis on the robustness of the localized active process across tissues of varying sizes and curvatures, particularly because the in vivo invagination is accompanied by global tissue growth and geometry changes.
Finally, we acknowledge that the simulated final shapes do not perfectly match the experimental geometry in every detail. We attribute these discrepancies to the omission of global tissue growth and the simplification of cell-cell adhesions between the surface epithelium and internal bulk cells. While these factors are not the primary drivers of the invagination, they undoubtedly refine the local morphology. We have added discussions of these limitations in the revised manuscript and aim to incorporate precise experimental measurements of tissue growth and inter-layer interactions in future modeling efforts.
Reviewer #3 (Public review):
Summary:
In this manuscript by Qiao et al., the authors seek to uncover force and contractility dynamics that drive tissue morphogenesis, using the Ciona atrial siphon primordium as a model. Specifically, the authors perform a detailed examination of epithelial folding dynamics. Generally, the authors' claims were supported by their data, and the conceptual advances may have broader implications for other epithelial morphogenesis processes in other systems.
Thank you for your positive summary and for recognizing the broader implications of our work.
Strengths:
The strengths of this manuscript include the variety of experimental and theoretical methods, including generally rigorous imaging and quantitative analyses of actomyosin dynamics during this epithelial folding process, and the derivation of a mathematical model based on their empirical data, which they perturb in order to gain novel insights into the process of epithelial morphogenesis.
Thank you for highlighting the strengths of our multidisciplinary methodology.
Weaknesses:
There are concerns related to wording and interpretations of results, as well as some missing descriptions and details regarding experimental methods.
We have revised the manuscript to address your concerns regarding the wording and the details of the methodology.
Recommendations for the authors:
Reviewing Editor Comments:
Based on the feedback from the reviewers, a focus on the following major points has the potential to improve the overall assessment of the significance of the findings and the strength of the evidence:
(1) It would be helpful to clearly articulate how these findings advance the field beyond what has already been demonstrated or suggested in other systems.
We thank the editor for this helpful suggestion. To better articulate how our findings advance the field, we have revised both the Introduction and Discussion to explicitly contrast our system with previously studied invagination models. Specifically, we highlight that our work demonstrates a bidirectional redistribution of actomyosin between apical and lateral domains, which differs from the apical-to-basolateral shift reported in ascidian endoderm invagination. Moreover, we emphasize that lateral contractility is obligatory for the accelerated invagination stage in our system, whereas in Drosophila ventral furrow invagination the second folding phase can proceed without it. These comparisons have been clearly presented in the revised manuscript. We think our findings represent a distinct mechanical paradigm for sequential epithelial morphogenesis.
(2) It would be helpful to clarify the meaning of "translocation" and more explicitly describe the temporal and spatial patterns of active myosin localization during the two steps of invagination.
We have replaced the term “translocation” with “redistribution” throughout the manuscript, including the title. We have also added new quantitative analyses of active myosin intensity from earlier time points (14–15 hpf) to rigorously support the initial lateral-to-apical redistribution phase (Figure 2B). High-resolution top-view images have been included to show the ring‑like localization of myosin at the apical cell‑cell junctions during the initial stage (Figure 2A). The schematic in Figure 2C has been corrected to accurately reflect the predominant localization of active myosin at the apical cell‑cell borders.
(3) It would be helpful to explain how the optogenetic data support the conclusion that "redistribution of myosin contractility from the apical to lateral regions is essential for the development of invagination".
We have performed additional experiments combining optogenetic inhibition with subsequent immunostaining of active myosin II (anti-pS19 MRLC). We quantitatively compared the distribution of actomyosin in light‑stimulated versus dark‑control embryos. The new data show that after light exposure, lateral myosin intensity is significantly reduced compared to the dark control, whereas apical myosin levels decrease similarly in both groups. This indicates that the optogenetic manipulation effectively attenuates lateral contractility during the accelerated invagination stage without affecting concurrent apical contractility changes. These results directly support the conclusion that lateral contractility acquisition is essential for invagination progression. (Figure 4—figure supplement 2)
(4) It would be helpful to describe how the modeling work fits within the existing literature on modeling epithelial folding and to address discrepancies between the model and the actual biological observations, such as tissue curvature, limited invagination depth in the model, and the "puckering" surrounding the invagination. In addition, certain descriptions of the modeling results should be clarified, as suggested by Reviewer #3.
We thank the referees for the detailed and constructive comments on our modeling work. In response to these suggestions, we have significantly updated the theoretical section of the manuscript. Specifically, we have reformulated the vertex model within a curved geometry that represents the entire tissue, and revised the subsequent analyses to better clarify the mechanical principles driving the observed morphogenesis. We have added relevant references and discussed the mechanistic connections and distinctions between our model and previous studies on epithelial invagination. We hope that our point-by-point responses of the modeling work and the corresponding revisions in the manuscript adequately address the reviewers’ concerns.
(5) It would be helpful to elaborate on the methods for quantitative image analysis and statistical tests.
We have thoroughly expanded the Materials and Methods section by adding a dedicated subsection “Quantification and statistical analysis”. This subsection provides step‑by‑step descriptions of how apical, lateral, and basal domains were defined (segmented line, width 1 μm), how normalization was performed (basal intensity set to 1), how center cell height, invagination depth, and lateral cell distance were measured (referencing Figure 1B), and what statistical tests were used (two‑tailed Student’s t‑test, with significance levels indicated). (see revised Materials and Methods, “Quantification and statistical analysis” subsection)
Reviewer #1 (Recommendations for the authors):
(1) This reviewer has concerns about two aspects of the model. First, the model in Figure 5D shows a simulation of a flat epithelial sheet creating an invagination. However, the actual invagination is occurring in a small embryo that has very significant curvature, such that nine or so cells occupy a 90-degree arc of the 360-degree circle that defines the embryo's section (e.g., see Figure 1A). This curvature could potentially have important effects on cell behavior. Ideally, the developed model would reflect the actual geometry of the observed behavior. A more nuanced analysis would provide important insight into whether the embryo's curvature makes a difference. Importantly, any result comparing the planar versus curved system would be interesting because if the model worked equally well in the high curvature or planar systems, the model is robust, or if invagination requires different strategies for high curvature and for planar systems, this is an important finding that reveals the importance of local geometries. I don't think the consideration of invagination from a planar vs curved epithelium has been previously modeled.
We fully agree with the reviewer that comparing planar versus curved systems provides valuable insights into the invagination mechanism. As we addressed in our response to Reviewer #1 (Public Review) - Weakness (1), we have now updated our vertex model to incorporate curved geometries and introduced surface bending stiffness to better reflect the embryo's actual shape. Our systematic comparison reveals that the invagination process, driven by apico-basal tension imbalance and lateral contraction, is indeed highly localized and remains robust across different initial curvatures. We have added Figure 5—figure supplement 1 and corresponding discussions in the revised manuscript to highlight these findings on model robustness and the role of local geometry.
(2) The second concern about the model is that Figure 5D shows the vertex model developing significant "puckering" (evagination) surrounding the invagination. Such "puckering" is not seen in the in vivo invagination (Figures 1A, 2A). This issue is not discussed in the text, so it is unclear how big an issue this is for the developed model. A discussion of this issue in the text would be appropriate. Maybe puckering goes away if a curved epithelium is modeled?
Thank you for this comment. In our model, the "puckering" effect naturally arises due to the presence of surface bending stiffness and the absence of rigid boundary constraints, which resembles the tissue morphology observed at 17 hpf in our experiments. However, our updated simulations show that this effect significantly diminishes as the tissue curvature decreases. We have addressed this concern in detail in our response to Reviewer #1 (Public Review) - Weakness (2) and have included the relevant analysis and discussions in the revised manuscript.
(3) Because of the puckering, it is unclear in the model what measurement is being used to define the invagination depth in Figure 5E. Is the depth from the maximal height of the surrounding epithelial cells? Or the location of the apical surface before invagination begins? It would be helpful to have that parameter better defined, and it would also be helpful to add a line to Figure 5D showing how the reference point for invagination depth.
Thank you for your suggestion. We measured the vertical distance from the baseline connecting the maximal height of apical midpoints of the surrounding cells to the apical surface of the center cell, which is consistent with our experimental measurements. We have now added a schematic line and indicators to Figure 5D.
(4) In Figure 2A Top View, as well as the schematic in Figure 2C, the developing invagination is surrounded by a ring of aligned cell edges characteristic of a "purse string" type actomyosin cable that would create pressure on the invaginating cells, which have been documented in multiple systems. Notably, the schematic in Figure 2C shows myosin II localizing to aligned "purse string" edges, suggesting the purse string is actively compressing the more central cells. If the purse string consistently appears during siphon invagination, a complete understanding of siphon invagination will require understanding the contributions of the purse string to the invagination process. For this paper, a discussion of the possible involvement of a purse string would be helpful for the readers, but follow-up work could include laser cutting or optogenetic blockage of the purse string contractility.
Thank you for your suggestion. We agree that the ring-like actomyosin structure is a prominent feature during the initial stages of invagination, and its potential role warrants discussion. We carefully re-examined our data. Our analysis confirms that this myosin ring is most pronounced during the early initial invagination stage (Figure 2A). This inward compression from the periphery would work in concert with apical constriction to help shape the initial invagination. However, this ring-like myosin pattern significantly diminishes in the accelerated invagination stage. We propose that the purse string may play a collaborative role in the early phase. We agree that follow‑up work (e.g., laser cutting or optogenetic manipulation) would be valuable and have noted this as a future direction in the Discussion.
(5) The introduction and discussion put the work in the context of work on physical forces in invagination, but there is not much discussion of how the modeling fits into the literature. Did the current work advance the state of modeling of such phenomena? What were the strengths and limitations of the modeling in this paper compared to what has been done previously?
Thank you for this suggestion. While we have incorporated additional literature in the revised manuscript as mentioned in our response to Reviewer #1 (Public Review) - Weakness (4), we would like to further clarify the specific advances and limitations of our modeling framework. Our updated vertex model builds upon established foundational frameworks but advances the state of modeling by: (i) incorporating dynamic apico-lateral tension variations coupled with actomyosin signals, and (ii) achieving localized, activity-mediated morphogenesis without the need for external rigid boundary constraints—a feature that distinguishes it from many classical models. We also recognize the model's current limitations. Specifically, it does not explicitly account for compressive stress and global geometric changes induced by tissue growth. The mechanical interactions between surface epithelial cells and the underlying internal bulk cells are also simplified. These factors represent important directions for our future work. We have added a dedicated paragraph in the Modeling and Discussion sections to contrast our model with existing literature and to explicitly state these strengths and limitations.
(6) Figure 4D. Minor point, but the labeling on the X-axis is out of register with the bar graphs.
We have corrected the alignment of the X‑axis labels with the bar graphs in Figure 4D. The figure has been updated accordingly.
(7) Figure 4B does not have a scale bar.
We have added a scale bar to Figure 4B (10 μm).
Reviewer #2 (Recommendations for the authors):
(1) Live imaging is necessary to demonstrate bidirectional translocation by visualizing the movement of the actomyosin network between the apical and lateral domains. Alternatively, a term other than "translocation" should be used to describe the observation.
We agree that live imaging of actomyosin movement would be ideal but is technically challenging in this system. Instead, we have replaced the term “translocation” with the more accurate and conservative term “redistribution” throughout the manuscript, including the title, to avoid implying physical movement of the same molecules. This addresses the reviewer’s concern.
(2) The optogenetic tool could be used to its full potential by manipulating myosin spatially or temporally, for example, by inhibiting myosin at various stages or subcellular locations, which would provide an opportunity to thoroughly test the domain and stage-specific needs for actomyosin. That said, I recognize that such experiments may be challenging in the model system used in this study.
We thank the reviewer for this suggestion. We have indeed attempted spatially restricted optogenetic activation in the Ciona atrial siphon system, but found it technically very challenging due to tissue geometry and light scattering. We appreciate the reviewer's understanding of these technical limitations.
(3) Some additional characterization of the optogenetics tool, such as the distribution of active myosin and F-actin post-stimulation, could further strengthen the interpretation of the inhibitory effect on invagination.
We thank the reviewer for this suggestion. After optogenetic inhibition, we fixed and stained embryos for active myosin II. The results (Figure 4—figure supplement 2) show that light exposure significantly reduces lateral myosin intensity compared to the dark control, while apical myosin decreases similarly in both groups. This confirms that the optogenetic manipulation selectively attenuates lateral contractility without affecting apical changes. We have added this data to the Results section.
(4) It would be helpful to address how heterogeneity in MRLC mutant overexpression might impact the interpretation of the outcome.
We acknowledge that mosaic expression is common in Ciona electroporation. For all quantitative analyses, we only selected embryos in which the center cell and more than half of the surrounding cells in the primordium showed clear expression of the plasmid. This selection criterion has been added to the Materials and Methods section.
(5) For Figure 2, it would be helpful to include the en face view of the cells at different apical-basal depths to better demonstrate the changes in the subcellular localization of myosin at different stages.
We have added top‑view images in Figure 2A at both the apical and a deeper (lateral) plane. These images clearly show the ring‑like localization of active myosin at the apical cell‑cell junctions during the initial stage. Together with the cross‑sectional views, they adequately demonstrate the subcellular localization changes.
(6) The Methods section should include more detailed descriptions of image quantification procedures. For example, for Figure 2B, how were the apical and lateral signals defined, and how were background intensities determined? In addition, the methods used for statistical tests should be clearly stated.
We agree that detailed quantification procedures are essential. We have therefore expanded the Materials and Methods with a new subsection “Quantification and statistical analysis”. This subsection includes precise definitions of apical, lateral, and basal domains (segmented line, width 1 μm), background subtraction (region outside the tissue), normalization (basal intensity set to 1), and descriptions of how cell height, invagination depth, and lateral distance were measured (referencing Figure 1B). Statistical tests (two‑tailed Student’s t‑test) and significance levels are clearly stated.
(7) The discrepancies between the model and experimental data, as described above, should be acknowledged. Commentary on how the model's assumptions and setup might contribute to these differences would be helpful.
We thank the reviewer for this suggestion. As detailed in our response to Reviewer #2 (Public Review) - Comment (6), we have included the discrepancies between the model and experimental results in the Modeling and Discussion sections. We have added comments explaining how our key modeling assumptions might contribute to these differences. Specifically, while we have updated the model to a curved geometry, the omission of continuous global tissue growth and expansion could affect the final invagination depth and shape. Meanwhile, the neglect of mechanical interactions between the surface epithelium and the internal bulk cells prevents the model from fully capturing the constraints that refine the local furrow configuration in vivo. By clarifying these limitations, we now provide a more balanced view of the model's scope and its role in identifying the primary mechanical drivers of invagination.
Reviewer #3 (Recommendations for the authors):
General comments:
(1) Methods: More information is needed to describe how imaging and quantification were performed. A couple of examples:
(a) In Figure 1, how were the apical and basal surface area of the center cell quantified?
(b) In Figure 1, Supplement 1, how was fluorescence intensity measured? Was there a constant area or volume that was quantified between samples? This is important because a decreasing apical surface can cause the signal to appear "concentrated" and increased.
We thank the reviewer for this important suggestion. We have added a dedicated subsection “Quantification and statistical analysis” in the Materials and Methods. This subsection includes precise definitions of apical, lateral, and basal domains (segmented line, width 1 μm), background subtraction (region outside the tissue), normalization (basal intensity set to 1), and descriptions of how cell height, invagination depth, and lateral distance were measured (referencing Figure 1B). Statistical tests (two‑tailed Student’s t‑test) and significance levels are also stated.
(2) The manuscript could use some editing and proofreading for grammar.
The manuscript has been carefully edited for grammar and clarity. We thank the reviewer for the suggestion.
Specific points:
(1) Figure 1A: Could the authors please annotate the location of the center cell throughout the time course? This would make it easier for the reader to understand what is being quantified.
We have added arrows to indicate the center cell at each time point in Figure 1A. This makes it easier for readers to follow the quantification.
(2) Figure 1 Supplement 1A, Line 143, "...before 15 hpf, F-actin concentration decreased at the lateral domains..."
It is not clear that the graph shows a decrease in the lateral domains when taking the error bars into account. It is possible that the F-actin concentration is stable in the lateral domains before 15 hpf. Are there some statistical analyses that can be performed?
We re-analyzed the F-actin data and agree that the change before 15 hpf is not statistically convincing given the error bars. However, we have added new quantitative analysis of active myosin (p-MLC) at 14–15 hpf (Figure 2B), which shows a clear and significant shift from lateral to apical enrichment during this early phase. This myosin dynamic strongly supports our hypothesis of bidirectional redistribution. The corresponding text has been updated in the Results section.
(3) Figure 1 Supplement 1A, Line 147-148, "...after 16 hpf, during which apical F-actin levels showed a gradual decline." Based on the graph, it does not appear that apical F-actin levels show a gradual decline after 16 hpf; rather, they may be steady or slightly increase.
We agree with the reviewer. Our original statement was inaccurate. What we intended to emphasize was that at 16 hpf, the F-actin level at the lateral domain exceeded that at the apical domain. The detailed changes of F-actin after 16 hpf were not a focus of our discussion. We have revised the text accordingly to avoid any misinterpretation. The correction has been made in the Results section.
(4) Figure 2C Hypothesis and line 169-170, "Initially, actomyosin translocated from the lateral regions to the apical domains..."
Related to the comment above, it is not clear that one can state that the actomyosin "translocated". The quantification does not necessarily demonstrate a loss of actin at the lateral domain in the initial stage, and even if there was a loss of lateral actomyosin, one would require experiments (perhaps photoconversion experiments) to demonstrate that machinery from the lateral region was transferred to the apical surface, rather than a process of new assembly at the apical surface.
We fully agree with the reviewer. We have replaced the term “translocation” with “redistribution” throughout the manuscript, including the title, to avoid implying physical movement of the same actomyosin structures. The text in the Results and Discussion has been revised accordingly.
(5) A similar comment is relevant to the subsequent statement in line 175, "actomyosin translocated from the apical domains to the lateral regions." Without direct experiments to demonstrate movement of the actomyosin machinery, it is possible that there is de novo assembly of actomyosin in the lateral region rather than translocation.
This wording ("translocation") becomes important primarily because it is in the title and appears to be one of the authors' major conclusions.
We fully agree with the reviewer that the wording is critical given our main conclusion. We have therefore systematically replaced “translocation” with “redistribution” across the manuscript (title, results, and discussion).
(6) Figure 4, Lines 215-216, "These results confirm that the redistribution of myosin contractility from the apical to lateral regions is essential for the development of invagination."
This experiment did not specifically test the redistribution of myosin; rather, the authors demonstrated that myosin contractility globally is necessary for invagination. In these experiments, is it known where the myosin is?
We have performed additional immunostaining experiments (new Figure 4—figure supplement 2) to directly examine myosin distribution after optogenetic inhibition. The results show that light exposure specifically reduces lateral myosin intensity compared to the dark control, while apical myosin decreases similarly in both groups. This demonstrates that the optogenetic manipulation selectively attenuates lateral contractility. We have revised the conclusion to state that the acquisition of lateral contractility is essential for invagination progression. The new data and revised text are in the Results section.
(7) Figure 4B, minor point: It would be helpful if the authors included a timestamp for the bottom row images (Dark 1 h).
Thank you for pointing out this typo. Timestamps have been added to the bottom row images (Dark 1 h) in Figure 4B.
(8) Figure 5E, F, minor point: It seems that the label on the red curve has a typo; it should be T18ES19E (rather than T18AS19E).
Thank you for pointing out this typo. We have corrected it in the revised manuscript (now Figure 6A, B).
(9) Figure 5F and corresponding text: Can the authors please clarify what is meant by "Coupled mode" as marked in the schematic? Is this meant to refer to simultaneous apical constriction and lateral contraction? Or sequential?
We thank the reviewer for this question. By "coupled mode," we refer to the mechanical synergy between apical and lateral contractions in driving the final invagination. As observed in our experimental data and recapitulated in the model, these two processes occur sequentially rather than simultaneously. We have revised the corresponding text to explicitly clarify this sequential process.
(10) Figure 6A, B, Lines 274-275: "...the invagination depth increased significantly under higher alphaa (Figure 6A), while the central height remained relatively independent of alphaa (Figure 6B)." This caused me some confusion until I realized that "Figure 6B" might be a typo and should be Figure 6C.
We sincerely apologize for this confusion. In the revised manuscript, this specific section and the corresponding figures have been updated.
(11) Line 287, typo: I believe that "Figure 5B" should be Figure 6B.
We sincerely apologize for this confusion. In the revised manuscript, this specific section and the corresponding figures have been updated.
(12) Figure 6A, B, comparing invagination depth with varying apical or lateral actomyosin intensity: The authors state that "invagination depth increased significantly under higher alphaa", but describe "mild invagination depth variation" with varied lateral actomyosin intensity. The graphs seem to suggest that there is increased invagination depth when either apical or lateral actomyosin intensity is increased, and that the increase is to a similar extent. Can the authors comment on what they think the differences are, if the apical effect is "significant" but the lateral effect is "mild"?
We thank the reviewer for this meticulous observation. We agree and feel sorry that our original description was not sufficiently precise. In the revised manuscript, we have re-analyzed the distinct contributions of apical and lateral tensions using the updated curved vertex model, which provides a more accurate mechanical decoupling. We have accordingly replaced the previous wording with a more rigorous description of the simulations and streamlined the corresponding figures to ensure the conclusions are clearly supported.
(13) Figure 6H, Lines 307-309, "...stronger regional translocation and redistribution contribute to the rapid reduction in height of invaginating cells..."
It appears from the graph that this is really only apparent at high alpha (total actomyosin); at empirically determined levels (alpha = 1), the effect of varying ratio is less dramatic. Can the authors comment on how significant they consider this effect?
We thank the reviewer for this insightful comment. We agree that the theoretical predictions regarding translocation strength in the original model lacked sufficient experimental validation. To maintain the scientific rigor of our study, we have removed the sections concerning the translocation ratio and the corresponding Figure 6H from the revised manuscript. Instead, we now refocus our analysis on the core mechanical drivers of invagination that are directly supported by our observations. We also have added discussions acknowledging other factors not fully captured in the current model (e.g., tissue growth), which we aim to investigate in future work.
