Regionalized regulation of actomyosin organization influences cardiomyocyte cell shape changes during chamber curvature formation

Domain: biology
Published: 2026-03-10
Editorial outcome: major_revision

Abstract
Cardiac chambers emerge from a heart tube that balloons and bends to create expanded ventricular and atrial structures, each containing a convex outer curvature (OC) and a recessed inner curvature (IC). The cellular and molecular mechanisms underlying the formation of these characteristic curvatures remain poorly understood. Here, we demonstrate in zebrafish that the initially similar populations of OC and IC ventricular cardiomyocytes diverge in the organization of their actomyosin cytoskeleton and subsequently acquire distinct OC and IC cell shapes. Altering actomyosin dynamics hinders cell shape changes in the OC, and mosaic analyses indicate that actomyosin regulates cardiomyocyte shape in a cell-autonomous manner. Additionally, both biomechanical cues and the transcription factor Tbx5a influence the basal enrichment of actomyosin and squamous cell morphologies in the OC. Together, our findings demonstrate that intrinsic and extrinsic factors intersect to control actomyosin organization in OC cardiomyocytes, which in turn promotes the cell shape changes that accompany curvature morphogenesis.

Introduction
Organs are replete with curvature, from the branching distal tips of the lung epithelium to the concave cup of the retina, and to the gyri and sulci of the brain. Essentially every organ exhibits some degree of curvature during its morphogenesis, and the precise curvature geometry required for a particular organ’s function is likely sculpted from inputs both extrinsic and intrinsic to its tissues 1. Decades of interest in this topic show us that some curvatures rely heavily on external influences. For example, the vertebrate gut acquires its dramatic twists and turns due to its rapid lengthening combined with its restrictive attachment to a relatively static partner, the dorsal mesentery 2. By contrast, the ingressing Drosophila ventral furrow is controlled largely by tissue-intrinsic processes, with apical constriction driven by actomyosin pulses in the epithelial cells resulting in robust invagination 3–5. Here, we investigate the factors, both tissue-extrinsic and -intrinsic, that control the regionalized cell shape changes that accompany chamber curvature formation in the developing heart.

The heart begins as a simple tube but then transforms into a looped organ with ventricular and atrial chambers. This looping process involves coordinated bending and ballooning of myocardial tissue to form the chambers, the twisting of the heart into a slightly helical shape (known as torsion), and the asymmetric displacement of the ventricle and atrium to deliver the chambers to their ultimate positions 6–10. During formation of the chambers, each acquires a bulging outer curvature (OC) and a recessed inner curvature (IC) 11–13. In addition to distinctive tissue morphology, these curvatures exhibit disparate physiological features; for example, the OC exhibits higher conduction velocity 14–16 and more trabeculation 17–19 than the IC, whereas the IC demonstrates greater tissue stiffness 20. While the differences between the OC and IC have been well characterized, the mechanisms that initially identify these regions and control their subsequent morphogenesis and specialization remain enigmatic.

Previous experimental and modeling studies have proposed cell behaviors such as hypertrophic growth, rearrangement within the tissue, and changes to cell shape as potential contributors to chamber curvature morphogenesis. For example, in chick, localized increases to cardiomyocyte volume in the OC appear to play a significant role in the creation of the ventricular bend 21,22. Additionally, in both chick and zebrafish, differing cell rearrangement patterns between the curvatures 9,23, as well as regionalized cell shape changes 24,25, likely play into the pronounced expansion of the OC tissue. In zebrafish in particular, the strikingly divergent morphologies that ventricular OC and IC cardiomyocytes attain during curvature formation have suggested that active cell shape change is a critical driver of curvature formation. Specifically, examination of the myocardial surface of the zebrafish heart tube has highlighted that its cardiomyocytes are fairly uniform in both apical surface area and circularity 25. However, after curvature formation, OC cardiomyocytes have a significantly larger, more elongated apical surface area than do IC cardiomyocytes 25. Consistent with the idea that regionalized cell shape changes create chamber curvatures, many mutations that cause chamber dysmorphia also disrupt the acquisition of OC- or IC-linked cell shapes 14,26–30. However, the subcellular mechanisms that drive these changes to cardiomyocyte morphology have not been well defined.

The actomyosin cytoskeleton is an attractive candidate for mediating cell shape changes. Although capable of inducing shape change in several ways, the network is perhaps best known for its ability to contract and thus create localized tension within a cell, as in the apical constriction observed during Drosophila ventral furrow formation 5,31. This type of actomyosin-derived tension is indeed employed at later stages of zebrafish cardiac morphogenesis, when differential tensions between cardiomyocytes are important for the decision to either trabeculate or remain in the compact layer 32. If actomyosin also plays an earlier role in curvature cell shape acquisition, we would expect to see different presentations of the network between curvatures. Indeed, in zebrafish, where there is a single layer of cardiomyocytes at this stage, visualization of the outer surface of the myocardium has highlighted what is interpreted as “cortical” F-actin in both curvatures (i.e. intense F-actin signal outlining each cardiomyocyte) and an additional pool of “cytoplasmic” F-actin running throughout OC cardiomyocytes 33. Although visualizations of the F-actin cytoskeleton in chick are somewhat more challenging due to the bilayered myocardium, distinctions between the OC and IC have been resolved, with circumferentially aligned F-actin at cell-cell boundaries in the IC and a less organized network of F-actin in the OC 34,35. Furthermore, actin dynamics appear to be necessary for the curvature formation process: in both chick and zebrafish, explanted hearts exposed to inhibitors of actin polymerization fail to achieve the stereotypic curved chamber contours 36–38. Altogether, these data position actomyosin as a likely regulator of curvature formation, but how the divergence of the actomyosin landscape is regulated and whether the network mediates its effect on tissue shape via active cell shape changes remain unknown.

Curvature formation takes place in a heart that actively beats and pumps blood, suggesting the possible involvement of biomechanical cues in regulating the cellular and subcellular changes that occur in OC and IC cardiomyocytes. It is well known that biomechanical inputs, such as fluid forces stemming from blood flowing through the endocardial lumen, tissue tension arising from the sarcomeric contraction of myocardial cells, and the anchoring of venous and arterial poles to the vascular system, influence several aspects of cardiac development 12,39–44. For example, altering the dynamics of blood flow in zebrafish can prevent both trabeculation 19,45,46 and valve formation 47–49. In addition, zebrafish hearts that lack atrial contractility and thus have reduced blood flow through the ventricle exhibit OC cardiomyocytes with a reduced apical surface area 25. This phenotype highlights the importance of extrinsic factors for attaining curvature-linked cell shapes, but it is not yet clear how these forces are translated into the subcellular events that instigate changes to cardiomyocyte morphology.

While extrinsic forces clearly impact curvature formation, factors intrinsic to the myocardium are certainly also involved in regulating the cytoskeleton, changing cell behavior, and ultimately sculpting tissue shape. In considering intrinsic factors that could influence curvature morphogenesis, the TBX family of transcription factors stand out as potentially important contributors. A set of these factors work in concert to pattern the expression of genes associated with the chamber curvatures: TBX5 promotes expression of OC-related genes, whereas TBX2 and TBX3 suppress expression of these genes 9,10,50–56. For example, the combination of these three factors determine the OC-enriched expression of Nppa in mice 51–54. In zebrafish, expression of nppa is already enriched in the future ventricular OC region of the linear heart tube, well before curvatures take shape 25. Although loss-of-function analyses indicate that nppa itself is not necessary for curvature formation 57, its early prepattern signifies that there may be other factors regulated by Tbx factors that could influence the ability of OC or IC cardiomyocytes to change their morphology. This idea is especially intriguing in light of recent work in mouse showing that TBX5 influences epithelial tension and cell morphology in the second heart field 58.

In this study, we delve into new aspects of how OC and IC cardiomyocyte morphologies diverge during curvature formation in zebrafish. Specifically, by studying OC and IC cardiomyocyte shapes in three dimensions, we have found that OC cardiomyocytes expand primarily in the planar axis, becoming more squamous as curvatures form, while IC cardiomyocytes extend primarily in the apicobasal axis, becoming more cuboidal. We show that these cellular changes are preceded by regionalized patterns of subcellular actomyosin organization, and that actomyosin plays a cell-intrinsic role in determining cardiomyocyte cell shape. Intriguingly, this role appears to be in the induction of planar spread, rather than the role of constriction for which the actomyosin cytoskeleton is so well known. Finally, we show that blood flow through the ventricle and tbx5a each promote the curvature-associated divergence of actomyosin organization and cardiomyocyte morphologies, with particularly prominent effects in the OC. From these findings, we propose that several extrinsic and intrinsic factors converge to control the cytoskeletal dynamics that govern cell shape changes, and that these changes to cardiomyocyte morphologies are a critical component in sculpting the characteristic contours of the cardiac chambers.

Results
Curvature formation coincides with the divergence of ventricular OC and IC cardiomyocyte morphologies

To study the cellular and molecular underpinnings of curvature formation, we first needed to develop a working method to standardize the boundaries of the ventricular curvatures. For this purpose, we used a combination of gene expression patterns and measurements of morphological features to guide boundary placement in the embryonic zebrafish ventricle at 48 hours post-fertilization (hpf) (Supplementary Fig. 1a-g). For gene expression, we relied on nppa, which is enriched in the OC of the ventricle 25,59. Generally, the highest nppa-expressing area of the ventricle was deemed to be the OC, and the lowest nppa-expressing area was deemed to be the IC (Supplementary Fig. 1a,b). After examining the nppa expression pattern in 12 wild-type ventricles at 48 hpf, we used the typical territory of nppa expression and its spatial relation to the atrioventricular canal (AVC) and distal end of the outflow tract (OFT) to determine the morphological features of the heart that could serve as boundaries for the curvatures (Supplementary Fig. 1e; see Methods for details). Once we had drawn these boundaries, we found that the IC region coincided satisfyingly with the expression of mb (Supplementary Fig. 1c,d), a gene previously noted to be enriched in the ventricular IC 60. In a wild-type heart at 48 hpf, the OC region defined by this method typically includes ~50-60 cardiomyocytes, and the IC region typically includes ~25-30 cardiomyocytes. We modified this method slightly to determine the curvature boundaries in wild-type hearts at 36 hpf (Supplementary Fig. 1h; see Methods for details), and we used these rules to bound the curvatures in mutant hearts as well (see subsequent figures, including Figs. 4-7). Although we think that these boundaries encompass the territories that are important for the study of ventricular curvature formation, it is possible that they also include some less relevant areas. Nevertheless, this method provides a useful and reproducible strategy for outlining the ventricular OC and IC in multiple contexts.

Our previous work has shown that certain characteristics of cardiomyocyte shape, such as apical surface area and circularity, are uniform throughout the myocardium at the linear heart tube stage (24 hpf), but eventually diverge in the ventricular curvatures by 48 hpf 25. Employing our new definitions of the OC and IC, we aimed to understand this divergence with greater temporal detail by looking at a developmental stage midway through curvature formation, in both the planar (X/Y) and apicobasal (Z) axes. At 37 hpf, when curvatures have just started to take shape, we found that OC and IC cardiomyocytes exhibit only a modest difference in morphologies (Fig. 1a-f,m-p). OC cardiomyocytes have a slightly but significantly larger apical surface area than IC cardiomyocytes, whereas the circularity of the apical surfaces are comparable (Fig. 1a-c,m,p). Additionally, cell thickness, as measured by apicobasal length, is similar between OC and IC cardiomyocytes at this stage (Fig. 1d-f,n). Both OC and IC cardiomyocytes increase in volume between 37 and 48 hpf and extend this new volume in both planar and apicobasal axes (Fig. 1g-l,o). However, OC cardiomyocytes expand along their planar axis to a greater extent than do IC cardiomyocytes (Fig. 1a-c,g-i), as observed by a greater increase in apical surface area (Fig. 1m), and they become more elongated, typically along their circumferential axis (Fig. 1g-i,p). By contrast, IC cardiomyocytes become taller along their apicobasal axis and remain more circular than OC cells (Fig. 1d-f,j-l,n,p). Together, these data suggest that cardiomyocytes in the developing curvatures grow to similar degrees during this time, but they choose to allocate their new volume along different axes: along the planar axis for OC cardiomyocytes and along the apicobasal axis for IC cardiomyocytes. As a consequence, OC cardiomyocytes become more squamous, and IC cardiomyocytes become more cuboidal. These data also place the onset of cardiomyocyte shape change at an early stage when the heart is still relatively tubular, highlighting active cell shape change as a potential driver of curvature formation.

Divergence of cardiomyocyte morphologies is preceded by changes in actomyosin organization

To understand the cellular mechanisms behind the divergence in OC and IC cardiomyocyte shapes, we first wanted to identify when previously reported cytoskeletal characteristics of the curvatures diverged 33–35. Specifically, do the portions of the ventricle that will become the IC and OC exhibit differential actomyosin landscapes as soon as the linear heart tube is formed, or do they diverge sometime during curvature formation? If the latter is true, does this divergence precede the appearance of the observed differences between OC and IC cell shapes? Throughout the process of curvature formation, we found that F-actin primarily resides along the inner surface of the cardiomyocyte membranes. At 28 hpf, when a small bulge of OC is just becoming evident (Supplementary Fig. 3a), subcellular distribution of F-actin appears fairly uniform in the presumptive OC and IC (Supplementary Fig. 3b-d). As observed previously 63, proximal ventricular cardiomyocytes (those closer to the AVC) exhibit F-actin mostly at their basal membrane (Supplementary Fig. 3d), whereas distal ventricular cardiomyocytes (those closer to the OFT) exhibit F-actin distributed around all of the surfaces of their membrane, with particularly strong presence at the apical surface (Supplementary Fig. 3c). At 36 hpf, when OC and IC cardiomyocytes have begun to show small morphological differences (Fig. 1a-f,m-p and Supplementary Fig. 3e), the F-actin in OC cardiomyocytes remains more enriched at the basal surface, particularly in the proximal ventricle (Supplementary Fig. 3f-h). In IC cells, F-actin is distributed more equally around the membrane surfaces, with a substantial amount at the lateral and apical surfaces (Supplementary Fig. 3f-h). At 50 hpf, OC and IC cardiomyocytes look fairly similar to each other, with F-actin distributed around all membrane surfaces and particularly intense signal at the lateral membranes (Supplementary Fig. 3i-l). Our observations that OC and IC cardiomyocytes begin with similar F-actin organization but diverge at 36 hpf, only to converge again by 50 hpf, drew our attention to 36 hpf as a stage worthy of more intense study. These findings correspond well with published visualizations of the myocardial surface showing that F-actin organization differs between OC and IC cells at 36 hpf (but not at 24 hpf) 33 and that myosin vb mutants, which fail to undergo OC cell planar expansion, exhibit aberrant F-actin organization in OC cells at 36 hpf 64.

To better understand these qualitative findings, we adopted a more quantitative approach to the examination of the subcellular localization of F-actin and phospho-Myosin (pMyosin) in individual cardiomyocytes of the OC and IC (Fig. 2a-e). For every cardiomyocyte analyzed, we quantified signal intensities to reveal what proportion of the total F-actin and pMyosin is localized to its basal, lateral, and apical membranes (Fig. 2b; see Methods for details). Using this approach, we found that OC cardiomyocytes have a higher proportion of basal actomyosin than do IC cardiomyocytes, whereas IC cardiomyocytes have a higher proportion of apical and lateral actomyosin than do OC cardiomyocytes (Fig. 2c-e). In addition, we wondered whether we could observe disparities between cardiomyocytes in the proximal OC/IC and cells in the distal OC/IC, as in Supplementary Fig. 3. Comparing these subsets of cells, we found that proximodistal differences are quite clear in the OC (Supplementary Figs. 4 and 5). In proximal OC cardiomyocytes, F-actin and pMyosin are highly enriched at the basal surface, whereas in the distal OC, cardiomyocytes have a significantly lower proportion of actomyosin at the basal membrane and a higher proportion of actomyosin at the lateral and apical membranes, giving an overall organization more similar to cardiomyocytes of the IC, which do not exhibit striking differences between proximal and distal regions. These data from individual cells (Fig. 2) support what we observed at the tissue level (Supplementary Fig. 3) and highlight a potentially important aspect of cytoskeletal dynamics during a critical period of curvature formation.

Altogether, our observations indicate that, in the linear heart tube, cardiomyocytes in the future OC and IC have similar actomyosin organization, and that the greater disparity lies between the proximal ventricle and the distal ventricle. As chambers begin to form, cardiomyocytes in the OC (particularly those in the proximal region) retain basal enrichment of actomyosin, while cardiomyocytes in the IC accumulate actomyosin at their apical and lateral surfaces. Notably, the process of acquiring these differential cytoskeletal landscapes divergence closely precedes the initiation of the cell shape changes observed in Fig. 1. These data suggest that the actomyosin network in cardiomyocytes is differentially regulated depending upon their location within the ventricle. In addition, these data imply potentially distinct roles for actomyosin in the curvatures, given that the precise location of actomyosin within a cell determines its ability to engage drivers of specific cell behaviors.

Modulation of actomyosin dynamics dampens the divergence of OC and IC cardiomyocyte morphologies

Given the intriguing regionalized patterns of subcellular actomyosin localization that arise during chamber curvature formation, we wondered whether the actomyosin cytoskeleton plays a role in the divergence of curvature cell shapes. Previous studies in chick and zebrafish have shown that actin polymerization is crucial for chamber curvature formation 36–38, but these studies did not delve into the impact of actin polymerization on cardiomyocyte morphology. We treated wild-type embryos from 24-36 hpf or from 36-48 hpf with either Latrunculin B (Lat B) or Blebbistatin (Bleb) to block actin polymerization or non-muscle myosin II (NMII) activity, respectively (Supplementary Fig. 6). In embryos treated with either LatB or Bleb from 24-36 hpf, OC cardiomyocytes exhibited significantly smaller apical surface areas when compared with DMSO-treated controls (Supplementary Fig. 6a,b,d,e,f,h). In contrast to the effects of treatment from 24-36 hpf, Lat B treatment from 36-48 hpf had no effect on the planar expansion of OC cardiomyocytes (Supplementary Fig. 6c,d), and, curiously, Bleb treatment from 36-48 hpf resulted in excessive planar expansion (Supplementary Fig. 6g,h). These data highlight 24-36 hpf as a particularly important window for actomyosin activity in promoting subsequent OC cardiomyocyte planar expansion. They also hint at a second phase of NMII activity between 36 and 48 hpf, during which Myosin-based contractility might be important for maintaining cardiomyocyte cell shape.

We next wanted to evaluate whether actomyosin activity is required in cardiomyocytes themselves, or if the effect we observed on cardiomyocyte shape upon LatB or Bleb treatment is due to a required function in another tissue, such as the endocardium. To assess these questions, we used established transgenic lines to express two versions of the non-muscle myosin light chain 9 gene (myl9) throughout the myocardium, driven by the cardiomyocyte-specific myl7 promoter, namely: wild-type myl9 (Tg(myl7:WT-myl9-mScarlet)) or a dominant negative version of myl9 (Tg(myl7:DN-myl9-eGFP)) 32. When assessing cardiomyocyte morphologies at 48 hpf, we found that expression of WT-myl9 resulted in cardiomyocytes morphologies reminiscent of those in wild-type embryos (Fig. 1g-l,m,n,p; Supplementary Fig. 7a-f,m-o). By contrast, expression of DN-myl9 resulted in OC cardiomyocytes with decreased apical surface area and increased apicobasal length when compared with OC cardiomyocytes expressing WT-myl9 (Supplementary Fig. 7g,h,j,k,m,n). Together, these changes resulted in a more cuboidal phenotype. Additionally, IC cardiomyocytes expressing DN-myl9 exhibited a slightly (but significantly) decreased apical surface area and equivalent apicobasal length when compared with those expressing WT-myl9 (Supplementary Fig. 7g,i,j,l,m,n). Thus, since myocardial expression of DN-myl9 results in OC and IC cardiomyocytes with relatively similar shapes, our data suggest that actomyosin dynamics are indeed important within the myocardium to support the divergence of OC and IC cell morphologies.

To better understand the effect of DN-myl9 expression on cardiomyocyte shape, we examined whether the changes in NMII activity that led to cell shape alterations at 48 hpf also had an effect on the F-actin network at 36 hpf. Immunostaining for pMyosin verified that the expression of DN-myl9 reduces NMII activity in the ventricular myocardium (Supplementary Fig. 8a-c), and we found a coincident reduction in myocardial F-actin in DN-myl9 ventricles (Supplementary Fig. 8a,b,d). By staining with DNase I 65–67, we found that this reduction in F-actin levels was not due to a reduction in available G-actin (Supplementary Fig. 8e-i). Together, these data confirm the efficacy of the Tg(myl7:DN-myl9-eGFP) transgene and suggest that NMII activity may be involved in polymerizing actin monomers into F-actin. We further found that subcellular localization of F-actin was disrupted in OC cardiomyocytes expressing DN-myl9 (Supplementary Fig. 9). Of particular interest, we found a shift in F-actin enrichment from the basal membrane to the apical and lateral membranes in DN-myl9-expressing OC cardiomyocytes (Supplementary Fig. 9a,b,e). This shift prompted us to quantify the ratio of the amount of F-actin at the basal membrane to the amount of F-actin at the apical and lateral membranes (basal / (apical + lateral)). This ratio revealed a striking shift away from the basal enrichment normally seen in WT-myl9-expressing OC cardiomyocytes (Supplementary Fig. 9f). From other contexts, we know that Myosin itself can alter F-actin characteristics. For example, it can support F-actin bundles by crosslinking 68–70, can induce disassembly of stress fibers 71–73, and can organize F-actin at specific locations within the cell 74–76. We posit that altering the function of the Myosin network by expression of DN-myl9 impacts both the abundance and organization of F-actin.

NMII function plays a cell-intrinsic role in promoting divergence of OC and IC cardiomyocyte morphologies

Having seen that cardiomyocytes fail to attain expected shapes when a dominant negative form of NMII is expressed throughout the myocardium (Supplementary Fig. 7), we were curious about whether actomyosin plays a cell-intrinsic role in promoting OC and IC cell shape divergence, or, rather, if NMII-driven tissue-level tensions are responsible for the acquisition of cell morphologies. We therefore performed blastomere transplantation to create genetically mosaic embryos (Fig. 3a). In these experiments, we analyzed hearts that contained donor-derived cardiomyocytes expressing DN-myl9 surrounded by host-derived wild-type cardiomyocytes (Fig. 3b-l; see Methods regarding how specific cardiomyocytes were chosen for analysis). In comparison with their wild-type host-derived neighbors, DN-myl9-expressing OC cardiomyocytes had a smaller apical surface area when compared with wild-type cardiomyocytes (Fig. 3b-e,j), but apicobasal length was unchanged (Fig. 3c,f,k). This OC cell phenotype is fairly similar to the effect of uniform expression of DN-myl9 throughout the myocardium (Supplementary Fig. 7g,h,k,m-o). To complement our blastomere transplantation approach, we also injected the Tg(myl7:DN-myl9-mScarlet) transgene plasmid into wild-type embryos at the one-cell stage and found reduced apical surface area in OC cardiomyocytes mosaically expressing the transgene when compared with neighboring cardiomyocytes with no detectable DN-myl9 expression (Supplementary Fig. 10).

In contrast to the effect of modulating NMII activity in OC cardiomyocytes, we found that DN-myl9-expressing donor-derived cardiomyocytes in the IC were very similar to their host-derived neighbors (Fig. 3g-l). As in the OC, this finding mirrors the effect of uniform expression of DN-myl9 throughout the myocardium (Supplementary Fig. 7g,i,l,m-o). Taken together, our analysis of cardiomyocytes expressing DN-myl9 emphasizes that actomyosin activity supports the divergence of OC and IC cell shapes by promoting the squamous identity of OC cardiomyocytes in a cell-intrinsic manner, perhaps by facilitating active expansion in the planar axis, as has been hypothesized 37. In contrast, IC cardiomyocytes appear to be comparatively unaffected by reduced NMII activity.

Divergence of OC and IC actomyosin organization and cardiomyocyte morphology depends upon biomechanical cues

Our results thus far had highlighted actomyosin organization and function as drivers of cell shape change during curvature formation, and we next wanted to know what types of upstream regulators might control these subcellular features. We were particularly curious to evaluate how blood flow might affect actomyosin localization and cardiomyocyte morphology, as our previous work has shown that myosin heavy chain 6 (myh6) mutants 77, which lack an atrium-specific myosin heavy chain and therefore lack atrial contractility, have reduced blood flow through the ventricle as well as reduced apical surface area of OC cardiomyocytes 25. Consistent with previous findings, we found that OC cardiomyocytes of myh6 mutants fail to extend along the planar axis (Fig. 4a,b,g,h,m; Supplementary Fig. 11). In addition, we observed that myh6 OC cardiomyocytes also extend inappropriately along the apicobasal axis (Fig. 4d,e,j,k,n; Supplementary Fig. 11). In contrast to the OC results, we found that the apical surface area and apicobasal length of myh6 IC cardiomyocytes are indistinguishable from those of wild-type IC cells (Fig. 4a,c,d,f,g,i,j,l-n; Supplementary Fig. 11); however, both OC and IC cells exhibit a modest but significant decrease in volume in myh6 mutants (Fig. 4o).

Given the effect of reduced blood flow on OC cell morphology, we next asked if this phenotype is coupled with a dysregulated actomyosin cytoskeleton. We found that the overall levels of pMyosin and F-actin in the ventricular myocardium in myh6 mutants at 36 hpf are reduced relative to wild-type (Supplementary Fig. 12a-d), similar to our observations in DN-myl9 transgenic hearts (Supplementary Fig. 8a-d). Also resembling the DN-myl9 scenario (Supplementary Fig. 8e-i), myh6 mutant hearts exhibit levels of G-actin similar to wild-type (Supplementary Fig. 12e-i), suggesting that the F-actin deficiency is due to a polymerization defect rather than limited availability of actin monomers. In addition to an overall deficit in F-actin, we found mislocalization of subcellular actomyosin in myh6 mutant OC cardiomyocytes (Fig. 5a-d,g,h): both F-actin and pMyosin shift away from the basal surface and to the lateral and apical surfaces in the proximal OC and distal OC, with a particularly striking shift in the proximal OC (Fig. 5c,d,g,h; Supplementary Fig. 12j). In contrast to this OC phenotype, both F-actin and pMyosin organization in myh6 mutant IC cardiomyocytes are comparable to wild-type (Fig. 5e-h; Supplementary Fig. 12j). Taken together, these data highlight a previously unappreciated role for extrinsic factors such as the biomechanical cues produced by blood flow in ensuring the regionalization of cytoskeletal landscapes and, subsequently, the planar expansion of OC cardiomyocytes.

Cell-autonomous function of tbx5a contributes to the divergence of OC and IC actomyosin organization and cardiomyocyte morphology

Alongside considering the extrinsic factors that influence curvature formation, we also wondered what types of intrinsic factors, operating within individual cells, could impact the divergence of cardiomyocyte cell shapes and actomyosin organization. We were particularly curious about the influence of the T-box transcription factor gene tbx5a. In addition to the known role for Tbx5 in promoting OC transcriptional programs 50,51,55, zebrafish homozygous for mutant alleles of tbx5a exhibit a highly dysmorphic heart by 48 hpf 9,78. As reported previously 78, the outward bulge of the tbx5a mutant OC is substantially reduced and the tbx5a mutant IC exhibits less of a kink when compared with a wild-type heart (Fig. 6a,g). We therefore investigated whether tbx5a mutant cardiomyocytes underwent the stereotypical divergence in shape associated with the ventricular curvatures. At the cellular scale, and similar to the phenotypes seen in DN-myl9-expressing hearts (Supplementary Fig. 7) and in myh6 mutants (Fig. 4), OC cardiomyocytes in tbx5a mutants fail to expand in the planar axis and undergo aberrant apicobasal expansion (Fig. 6a,b,d,e,g,h,j,k,m,n). This combination results in tbx5a mutant OC cardiomyocytes with a more cuboidal phenotype than wild-type OC cardiomyocytes (Supplementary Fig. 11). Interestingly, even though at the tissue level tbx5a mutant ICs fail to form the stereotypic kink (Fig. 6g), IC cell shapes in tbx5a mutants appear comparable to wild-type with just a slight trend toward reduction in apicobasal length (Fig. 6a,c,d,f,g,i,j,l-n; Supplementary Fig. 11). In contrast to myh6 mutants (Fig. 4o,p), tbx5a OC and IC cardiomyocytes exhibit volumes and circularity similar to that of wild-type cardiomyocytes (Fig. 6o,p).

We next wondered whether the effect of tbx5a loss-of-function on OC cardiomyocyte morphology was preceded by a dysregulation of the actomyosin cytoskeleton. Similar to the DN-myl9 (Supplementary Fig. 8a,b,d-i) and myh6 (Supplementary Fig. 12a,b,d-i) phenotypes, the overall levels of F-actin in the tbx5a mutant ventricular myocardium are reduced at 36 hpf, even though G-actin levels appear similar to that of wild-type (Supplementary Fig. 13a,b,d-i). In contrast to the DN-myl9 and myh6 phenotypes (Supplementary Figs. 8a-c and 12a-c), however, levels of pMyosin in tbx5a mutants are not significantly different from wild-type (Supplementary Fig. 13a-c), suggesting that Tbx5a-regulated pathways affect actin polymerization independently of NMII activity. In addition to an overall deficit in F-actin, we found mislocalization of subcellular F-actin and pMyosin in tbx5a mutants, and this was particularly prominent in the proximal OC (Fig. 7a-h). In the tbx5a mutant proximal OC, F-actin and pMyosin enrichment shift away from the basal surface and accumulate more at the lateral and apical surfaces (Fig. 7g,h; Supplementary Fig. 13j). In distal OC and IC cardiomyocytes, F-actin is also shifted laterally and apically (albeit to a lesser extent than in the proximal OC), but the ratio of basal to apical/lateral pMyosin is comparable to wild-type (Fig. 7g,h; Supplementary Fig. 13j). Intriguingly, in contrast to the myh6 mutant scenario where the effect on pMyosin localization is very similar to the effect on F-actin localization (Fig. 5g,h; Supplementary Fig. 12j), the F-actin phenotype observed in tbx5a mutants is more severe than the pMyosin phenotype (Fig. 7g,h; Supplementary Fig. 13j). Overall, these data suggest that Tbx5a-regulated pathways are particularly important for the enrichment of F-actin at the basal surface of OC cardiomyocytes.

Given the actomyosin and cell morphology phenotypes in flow-deficient myh6 mutants (Figs. 4,5) and the reduced heart rate in tbx5a mutants 78, we wondered whether the aberrant effects on F-actin localization in tbx5a mutants reflect indirect consequences of altered blood flow or a cell-intrinsic loss of tbx5a function. To address this, we took a mosaic approach in which we transplanted wild-type blastomeres into tbx5a mutant host embryos and tbx5a mutant blastomeres into wild-type host embryos, as well as performing wild-type into wild-type and mutant into mutant controls (Fig. 8a-d). We fixed host embryos at 36 hpf and assessed amounts of F-actin at basal, lateral, and apical membranes (Fig. 8e-p; Supplementary Fig. 14). In a tbx5a into wild-type scenario, we found that the F-actin organization in tbx5a mutant donor-derived proximal OC cardiomyocytes largely resembled that of their wild-type neighbors (Fig. 8k,m; Supplementary Fig. 14c), indicating that something about the wild-type environment can rescue this aspect of the tbx5a mutant phenotype. In light of this result, we were particularly intrigued by the wild-type into tbx5a scenario. Here, we found that wild-type donor-derived cardiomyocytes in a tbx5a mutant proximal OC contained a significantly higher proportion of basal F-actin compared with apical and lateral F-actin, when compared with their tbx5a mutant neighbors (Fig. 8h,j; Supplementary Fig. 14b). This suggests an ability of wild-type cardiomyocytes to organize their F-actin network regardless of the tbx5a status of surrounding cells. Together, these data indicate that there is at least a partially cell-autonomous role for tbx5a in establishing the divergent actomyosin organization observed in the developing curvatures, but that there are also likely to be non-autonomous ways that tbx5a influences actomyosin organization.

We next wanted to know whether tbx5a also plays a cell-autonomous role in promoting the divergence of OC and IC cardiomyocyte morphologies, or if the cardiomyocyte shape anomalies observed in Fig. 6 are due to cardiac function defects present in tbx5a mutants 78. To distinguish between these possibilities, we used the same experimental scheme as in Fig. 8, but we instead fixed host embryos at 48 hpf and assessed cardiomyocyte shapes (Supplementary Fig. 15). Here, we focused on assessing apical surface area, as our data suggest a primary role for actomyosin in promoting the planar expansion of cardiomyocytes, particularly in the OC. Interestingly, consistent with our analysis of actomyosin localization in wild-type into tbx5a mosaic hearts (Fig. 8h,j), wild-type donor-derived OC cardiomyocytes extended in the planar axis significantly more than their tbx5a mutant host-derived neighbors, allowing them to assume a more squamous identity (Supplementary Fig. 15h,j). Also like our mosaic analysis of actomyosin localization (Fig. 8k,m), tbx5a mutant donor-derived cardiomyocytes that resided in a wild-type OC largely exhibited morphology similar to their wild-type neighbors (Supplementary Fig. 15k,m). Finally, donor-derived IC cardiomyocytes in both wild-type into mutant and mutant into wild-type scenarios achieved morphologies similar to their host-derived neighbors (Supplementary Fig. 15i,j,l,m). Taking our transplantation experiments together, it appears that the environment of a wild-type heart can largely override the absence of tbx5a function in donor-derived mutant cardiomyocytes, but, conversely, wild-type cardiomyocytes can achieve stereotypical OC traits even in a tbx5a mutant background. This suggests that the ventricular phenotypes we see in tbx5a mutants are not solely due to defects in blood flow and that there is some cell-autonomous aspect to tbx5a function during curvature formation.

An unexpected finding from our mosaic studies was that, in some cases, donor-derived cardiomyocytes sent basal projections underneath their host-derived neighbors (Supplementary Fig. 16). We observed such projections in all four of our transplant scenarios; these projections were typically rather short and slim and were usually restricted to the junctions between neighboring cells (Supplementary Fig. 16b,e). However, in the wild-type into tbx5a mutant transplant scenario, it was not uncommon for wild-type donor-derived OC cardiomyocytes to send long, broad projections underneath the cell body of a neighboring tbx5a mutant host-derived cardiomyocyte (Supplementary Fig. 16c-e). This could be a reflection of the comparatively robust ability of wild-type cardiomyocytes to expand their basal domain, a relatively reduced cell-extracellular matrix (ECM) connection in tbx5a mutant tissue, or both. This idea is further supported by the observation that, in the same wild-type into tbx5a mutant scenario, donor-derived OC cardiomyocytes that did not contact any other donor-derived OC cardiomyocytes (i.e. those in single-cell clones) had, on average, a higher apical surface area than donor-derived OC cardiomyocytes that contacted other donor-derived cardiomyocytes (Supplementary Fig. 17a). Further morphological assessment of these single-cell clones showed that all of them sent out basal projections: 3 of the 7 clones sent thin basal projections that extended between the junctions of neighboring host cells (Supplementary Fig. 17b), and the remaining 4 sent larger basal projections underneath neighboring host-derived cells (Supplementary Fig. 17c). We propose that this ability to send basal projections is a mechanism that normally drives planar expansion of OC cardiomyocytes, and that this activity is increased when neighboring cardiomyocytes maintain a loose attachment to the underlying ECM. Additionally, these data further support a cell-autonomous role for tbx5a in directing cardiomyocyte behavior.

Methods
Zebrafish

The following transgenic lines were used in this study: Tg(myl7:WT-myl9-mScarlet)bns524Tg 32, Tg(myl7:DN-myl9-eGFP)bns333Tg 32, Tg(myl7:eGFP-Hsa.HRAS)s883Tg 61, and Tg(myl7:mKate-CAAX)sd11Tg 62. We also used lines carrying the tbx5am21 78 and myh6m58 77 mutations. Heterozygous carriers of mutations were identified by PCR amplification and subsequent restriction enzyme digest, as previously described for tbx5am21 30 and myh6m58 62. Homozygous mutant embryos were identified by previously characterized morphological phenotypes 77,78. All work presented here followed protocols (#S09125) approved by the Institutional Animal Care and Use Committee at the University of California, San Diego.

Pharmacological treatments

Embryos were dechorionated and placed in 6-well culture dishes and exposed to either 1.25% DMSO, 1.25% DMSO + 100 ng/mL Latrunculin B (Millipore Sigma, L5228), or 1.25% DMSO + 30 μM Blebbistatin (Millipore Sigma, B0560) in E3 medium. DMSO controls were kept in solution between 24-48 hpf. Latrunculin B and Blebbistatin treated embryos were either kept in drug-containing media between 24-36 hpf and switched to 1.25% DMSO between 36-48 hpf, or vice versa. At 48 hpf, all embryos were fixed and processed as described below.

Immunofluorescence and phalloidin and DNase I staining

Embryos were dechorionated, fixed in 1% methanol-free formaldehyde (ThermoFisher Scientific, 28908) for 1 hour and 10 minutes with gentle rocking, and then rinsed three times in PBT (1X PBS containing 0.1% Tween 20 (Sigma-Aldrich, P9416)). Embryos were rocked rapidly in a 0.2% saponin solution (Sigma-Aldrich, S4521) in PBT containing 0.5% Triton X-100 (G-Biosciences, 786513) until yolks were dissolved. This took 20-35 minutes and was monitored under a dissecting microscope. Embryos were rinsed three times in PBT, placed in 4% formaldehyde, and incubated at 4°C overnight. On the following day, to improve antibody signal-to-noise ratio, embryos were rinsed three times in PBT, placed in prechilled 100% acetone, and incubated at -20°C for 8 minutes. Acetone was replaced with PBT + 0.5% Triton X-100 for one hour at room temperature. To ensure reagent access to the heart, forceps were used to open the pericardial cavity. Embryos were then blocked with 2 mg/ml bovine serum albumin (Sigma-Aldrich, A9647) and 10% goat serum in PBT for at least one hour at room temperature, then incubated with primary antibody in block overnight at 4°C. Embryos were then washed extensively with PBT, blocked as described for primary antibody incubation, and incubated with the secondary antibodies in blocking solution overnight at 4°C. Embryos were then washed extensively with PBT.

The following primary antibodies were used at the specified dilutions: mouse anti-Alcama (Developmental Studies Hybridoma Bank, Zn-8 supernatant, 1:50); rabbit anti-Cdh2 (GeneTex, GTX125885, 1:200); rabbit anti-phospho-Myosin (Abcam, ab2480, 1:100); rabbit (Life Technologies, A11122, 1:500) or chicken (Life Technologies, A10262, 1:1000) anti-GFP; rabbit anti-dsRed (also detects mScarlet; Clontech, 632496, 1:1000); rabbit anti-TagRFP (also detects mKate; Evrogen, AB233, 1:500); mouse anti-myosin heavy chain (Developmental Studies Hybridoma Bank, MF20 supernatant, 1:50); and mouse anti-Myh6 (Developmental Studies Hybridoma Bank, S46 supernatant, 1:50).

All secondary antibodies were produced in goat by ThermoFisher Scientific and used at a dilution of 1:300. Secondary antibodies used were: anti-mouse-AlexaFluor 488 (A11001); anti-mouse-AlexaFluor 568 (A11031); anti-mouse-AlexaFluor 647 (A21236); anti-rabbit AlexaFluor 488 (A11008); anti-rabbit AlexaFluor 568 (A11011); anti-rabbit AlexaFluor 647 (A21245); and anti-chicken AlexaFluor 488 (A11039). In some cases, Phalloidin-AlexaFluor 488 or Rhodamine-Phalloidin (Life Technologies, A12379 or R415; each reconstituted to 200U/mL in MeOH) was added to the secondary antibody solution at a dilution of 1:30.

For Supplementary Figs. 8, 12, and 13, embryos were stained with DNase I-AlexaFluor 594 (ThermoFisher D12372; reconstituted to 5 mg/mL in phosphate-buffered saline, pH 7.4 (PBS) + 50% v/v glycerol), diluted 1:500 to detect G-actin 65,66, and Phalloidin-AlexaFluor 647 (ThermoFisher A22287; reconstituted to 66 µM in DMSO), diluted 1:300 to detect F-actin. Staining was performed similarly to immunostaining, but without the addition of Triton X-100 to any solutions to prevent extraction of G-actin 67 and without a blocking step. Although Phalloidin signal was robust in both myocardium and endocardium, DNase I signal was detected at substantially higher levels in the myocardium when compared with the endocardium, supporting the specificity of this reagent.

Fluorescent in situ hybridization

Fluorescent in situ hybridization was performed as previously described 99, but embryos were fixed and deyolked as for immunofluorescence before MeOH dehydration and were incubated with Proteinase K (50 μg/mL) in PBT for 2 minutes at room temperature prior to hybridization with probe. Probe for nppa (ZDB-GENE-030131-95) was produced as previously described 77. Probe for mb (ZDB-GENE-040426-1430) was produced by PCR amplification of 48 hpf embryonic cDNA followed by T7 in vitro transcription. The forward primer used was 5′-GGACAAACACCGCGACAGAC-3′ and the reverse primer used was: 5′-TAATACGACTCACTATAGGGCCTGAGACCCTAACGAACCATTAT-3′; underlined portion is the T7 binding site.

Transplantation

Blastomere transplantation was performed similarly to a previous description 25. Briefly, donor embryos were injected with 5% lysine-fixable rhodamine dextran (ThermoFisher Scientific, D1817) in 0.2M KCl at the one-cell stage. Approximately 10-30 blastomeres were taken from dechorionated 3-4 hpf donor embryos and placed into the margin of similarly staged host embryos. Host embryos were raised in E3 medium containing penicillin-streptomycin (Millipore Sigma, P4458; diluted 1:50), in agarose-coated wells. Host embryos were allowed to develop until 36 or 48 hpf when they were screened for the presence of donor-derived cells in the myocardium, which could be identified by fluorescence under a Zeiss Axiozoom microscope. Hosts containing donor-derived cardiomyocytes were fixed and processed for immunostaining as described above. In order to be included for analysis, cardiomyocytes needed to meet specific criteria. For donor-derived cells, we analyzed any cell with at least half of its body within the OC or IC boundary. For host-derived cells, we analyzed any cell (1) in direct contact with a donor-derived cell or one cell-distance from a donor-derived cell and (2) with at least half of its body within the OC or IC boundary.

For Tg(myl7:DN-myl9-eGFP) transplants, transgenic animals were crossed to wild-type animals, and the resulting embryos served as donors; wild-type embryos served as hosts. For the tbx5a transplantation experiments, animals heterozygous for the tbx5a mutation were crossed to obtain both donor and host embryos. After transplantations were complete, donor embryos were genotyped by PCR and restriction enzyme digest for the tbx5a mutation, as described above. Genotypes of host embryos were determined based on the presence or absence of pectoral fin buds at 36 hpf or pectoral fins at 48 hpf 78. While it is possible that a large donor-derived clone could affect the development of one pectoral fin bud, it is unlikely, given the number of donor cells transplanted, that this would impact both pectoral fin fields in the same host. Both sides were therefore assessed for the presence of pectoral fin buds/fins, and only those host embryos that had matching sides were analyzed.

Transgene plasmid injection

To obtain the mosaic expression of Tg(myl7:DN-myl9-mScarlet) 32 shown in Supplementary Fig. 10, a mixture of transgene plasmid (10-20 pg per embryo) and tol2 mRNA (20-60 pg per embryo) was injected into the yolk of wild-type embryos at the one-cell stage. Embryos were screened for mScarlet fluorescence at 48 hpf, then were fixed and immunostained for Alcama as described above.

Image acquisition

After all immunostaining and fluorescent in situ hybridization procedures, hearts were dissected from embryos under a Zeiss AxioZoom microscope. To keep organ morphology as intact as possible, hearts were simply mounted in a droplet of PBT on a cover slip and immediately imaged using a Leica SP8 laser-scanning confocal microscope and a 25X water-immersion objective. Hearts were positioned laterally, with one side of the ventricle flat against the coverslip. In all experiments where signal intensity was measured, laser settings were kept consistent between samples. Z-stacks were captured with a slice thickness of 0.3 μm.

Image analysis

All image files captured by the Leica SP8 confocal were rendered as 3D reconstructions in Imaris v11.0.0 (Bitplane). Often, secondary analysis was completed in FIJI v2.16.0 (ImageJ).

Defining the ventricular OC and IC

To develop a method to standardize the boundaries of the OC and IC between 48 hpf samples (Supplementary Fig. 1e), we first assessed the expression domain of nppa in 12 wild-type hearts at 48 hpf (Supplementary Fig. 1a,b). By using the following measurements, we found that we could capture, on average, the highest nppa signal in the resulting “OC” and, on average, the lowest nppa signal in the resulting “IC”. These territories also corresponded well to the regions with the lowest (OC) and highest (IC) mb expression (Supplementary Fig. 1c,d). These measurements were then applied to 48 hpf hearts throughout this study.

The following abbreviations are used in our description of this measurement method: PBOC: OC proximal boundary; DBOC: OC distal boundary; DBOC(OFT): distal boundary of the OC side of the OFT; PBIC: IC proximal boundary; DBIC: IC distal boundary; DBIC(OFT): distal boundary of the IC side of the OFT.

To delineate the OC in Imaris (Supplementary Fig. 1e), the arc of the ventricular OC was measured beginning one cell diameter from the AVC (the PBOC) to the DBOC(OFT). This arc is represented by a blue dotted line in Supplementary Fig. 1e. Starting from the PBOC, the DBOC was marked at 2/3 of the distance from the PBOC to the DBOC(OFT). An Oblique Slicer was placed between the PBOC and DBOC, ensuring that it was perpendicular to the arc of the OC. The position of the Oblique Slicer is represented by a solid blue line in Fig. S1e. The portion of tissue bounded by the solid blue line was considered the OC.

To delineate the IC in Imaris (Supplementary Fig. 1e), the arc of the ventricular IC was measured from the AVC (the PBIC) to the DBIC(OFT). This arc is represented by a dotted pink line in Supplementary Fig. 1e. Point I (red “I” in Supplementary Fig. 1e) was marked at 1/2 of the distance between PBOC and DBOC. Point II (red “II” in Supplementary Fig. 1e) was marked at 1/3 of the distance between PBOC and Point I. The proximal border of the IC was considered to run from the PBIC to Point II. This border is represented by a solid pink line in Supplementary Fig. 1e. Starting from the PBIC, the DBIC was marked at 1/2 of the distance from the PBIC to the DBIC(OFT). Point III (red “III” in Supplementary Fig. 1e) was marked halfway between the DBIC and DBOC. The distal border of the IC was considered to run from the DBIC to Point III. This border is also represented by a solid pink line in Supplementary Fig. 1e. Finally, an Oblique Slicer was placed between Points II and III, ensuring that it was perpendicular to the arc of the IC. The position of this Oblique Slicer is represented by a solid pink line in Supplementary Fig. 1e. The portion of tissue bounded by the solid pink lines was considered the IC.

To account for the reduced contours of the OC and IC in hearts at 36-37 hpf (Supplementary Fig. 1h), slight adjustments were made in our measurement method. Specifically, the arc of the ventricular OC was measured beginning one cell diameter from the AVC (the PBOC) to the DBOC(OFT). This arc is represented by a dotted blue line in Supplementary Fig. 1h. Starting from the PBOC, the DBOC was marked at 2/3 of the distance from the PBOC to the DBOC(OFT). The arc of the ventricular IC was measured from the AVC (the PBIC) to the DBIC(OFT). This arc is represented by a dotted pink line in Supplementary Fig. 1h. A line was drawn between the PBOC and the PBIC, as well as between the DBOC and the DBIC; these are represented by dotted purple lines in Supplementary Fig. 1h. Point I (red “I” in Supplementary Fig. 1h) was marked at 1/3 of the distance between PBOC and PBIC, and Point II (red “II” in Supplementary Fig. 1h) was marked at 2/3 of the distance between PBOC and PBIC. Point III (red “III” in Supplementary Fig. 1h) was marked at 1/3 of the distance between DBOC and DBIC, and Point IV (red “IV” in Supplementary Fig. 1h) was marked at 2/3 of the distance between DBOC and DBIC. The medial border of the OC was considered to run from Point I to Point III (solid blue line in Supplementary Fig. 1h), and the medial border of the IC was considered to run from Point II to Point IV (solid pink line in Supplementary Fig. 1h). As for 48 hpf hearts, Oblique Slicers were used in accordance with the medial border measurements in Supplementary Fig. 1h to bound the OC and IC.

Labeling lateral cell membranes

When the labeling of lateral membranes was necessary, we utilized one of four reagents: Tg(myl7:mKate-CAAX), Tg(myl7:Ras-eGFP), anti-Cdh2 antibody, or anti-Alcama antibody. The first two reagents label all membranes of the cardiomyocyte, whereas the latter two are cell adhesion molecules and are therefore expected to label only the lateral membranes. Given the possibility that Cdh2 and Alcama could be polarized along the apicobasal axis, we tested the extent to which these signals overlapped with Tg(myl7:Ras-eGFP) signal (Supplementary Fig. 2). The consistent overlap of Cdh2 and Alcama signal with lateral Ras-eGFP signal (Supplementary Fig. 2; 100) led us to consider both Cdh2 and Alcama as sufficient representations of the entire lateral membrane, and we therefore used antibodies against either, depending on the specific requirements of an experiment.

Cardiomyocyte morphometrics

For each cardiomyocyte in the OC and IC, Imaris was used to capture Snapshots of the necessary views of the cell, FIJI was used to take measurements, and Excel was used to compile these measurements into a .csv and to calculate apicobasal length and volume.

In Imaris: To capture the en face view to obtain apical surface area and circularity, a single Snapshot of the apical surface of the cardiomyocyte was taken, ensuring that the surface was flat and parallel to the screen. If a cardiomyocyte was large enough that it curved in the Z-plane, multiple Snapshots were captured. To do this, an Oblique Slicer (or more, if necessary) was placed where the greatest curvature existed, perpendicular to the long axis of the cell. Then two (or more, if necessary) Snapshots of the cell were taken — the first on one side of the Oblique Slicer, and a second on the other side of the Oblique Slicer, ensuring that each “side” of the cell was flat and parallel to the screen. To capture the cross-sectional view to obtain apicobasal length and volume measurements, two Oblique Slicers were placed while the cell was still in an en face view. These Oblique Slicers were perpendicular to the apical surface of the cell and approximately bisected the cell (one bisected the major axis and the other bisected the minor axis). The Volume of the 3D reconstruction was clicked Off, and the image was reoriented so that the “faces” of the Oblique Slicers were visible, showing cross-sections of the cell. The intersection of the two Oblique Slicers marked the current cell of interest. Two Snapshots of the cell were taken, one from each Oblique Slicer, ensuring that the Slicers were flat and parallel to the screen.

In FIJI: The scale was first set based on the scale bar provided by Imaris. To measure apical surface area and circularity, the Freehand Selection tool was used to trace the lateral membrane of the cardiomyocyte. If a cardiomyocyte was curved in the Z-plane and required two (or more, if necessary) Snapshots, the lateral membrane of one “side” of the cell was traced, stopping where the cell meets the Oblique Slicer. The selection was copied and pasted onto the Snapshot of the other “side”, the “sides” were positioned so that they joined together, and the border of the entire cell was traced. The Measure function was used to obtain Area and Circularity values. For each of the two cross-sectional Snapshots taken to measure apicobasal length, the basal width (the distance between the basal tips of the lateral membranes), apical width (the distance between the apical tips of the lateral membranes), and length of each lateral membrane (from basal tip to apical tip) were measured using the Measure function.

In Excel: To obtain apicobasal length, the average (mean) of the lengths of the four lateral membranes was calculated. To obtain volume, the average (mean) of the two basal width values and the two apical width values was calculated; then, the mean basal width, the mean apical width, and the mean apicobasal length were multiplied.

Subcellular actomyosin localization

Prior to image analysis, F-actin and pMyosin (where appropriate) signal in each ventricle was assessed in Imaris. Briefly, a Mask of the ventricular myocardium was generated, the Mean signal intensity for each channel was located in the Statistics tab, and these values were tested for outliers in R 101. Outliers were removed from further analysis.

For each cardiomyocyte in the OC and IC, Imaris was used to capture Snapshots of the necessary views of the cell, FIJI was used to take signal intensity measurements, and Excel was used to compile these measurements into a .csv and to calculate mean signal intensities.

In Imaris: Two cross-sectional views (Fig. 2a,b) were captured as described above in the “Cardiomyocyte morphometrics” section, with the exception that a separate Snapshot was taken for each channel (for the membrane marker, for F-actin, and for pMyosin).

In FIJI: All images were converted to 8-bit and the channels for each Snapshot were Merged. Using the membrane marker channel as a guide, the Freehand Line tool, set to a width of ~1 μm, was used to trace just inside the membrane. The Mean signal intensities of the F-actin signal and the pMyosin signal were measured along each membrane using the Measure function, toggling between channels as needed. Four Mean measurements were taken per Snapshot: basal, apical, and two lateral. To prevent the inclusion of signal from the lateral membranes into the basal/apical datasets and vice versa, a ~1 μm segment in each “corner” (where the lateral and basal/apical membranes intersect) was avoided when performing the initial traces.

In Excel: For F-actin and for pMyosin (when present) in each cell, the averages (means) of the two basal signal intensities, of the two apical signal intensities, and of the four lateral signal intensities were calculated.

Total pMyosin, F-actin, and G-actin in the ventricular myocardium

All analysis was completed in Imaris v11.0.0. A Surface of the myocardium was generated using a Surfaces Detail of 0.9 μm and Machine Learning segmentation to include the ventricular myocardium and to exclude the endocardium as well as the space around the myocardium. The Surface was Cut at the AVC/ventricle boundary and at the ventricle/OFT boundary. The Mean signal intensity of each channel was located in the Statistics tab. Finally, each experimental replicate was normalized to a scale of 1-10 relative fluorescence units (rfu) using Min-Max Normalization. This transformation allows for comparison of experimental replicates performed on different days while preserving the relationship between datapoints.

Comparing morphologies of OC and IC cardiomyocytes with different genotypes

In Supplementary Fig. 11, we sought to compare cardiomyocyte morphologies of myh6 and tbx5a mutants to wild-type, all on one scale. To accomplish this comparison, we normalized the experimental mutant data using the formula:

(1) mean norm.exp = mean ref + (SD ref * mean(Z-score exp)),

where the mean ref is the mean of the 48 hpf wild-type reference dataset presented in Fig. 1, the SD ref is the standard deviation of the 48 hpf wild-type reference dataset presented in Fig. 1, and the mean(Z-score exp) is the mean Z-score of the experimental group (from the data presented in either Fig. 4 or Fig. 6). For example, the mean apical surface area for the 48 hpf wild-type OC reference dataset in Fig. 1 is 142 µm2, and the standard deviation for this dataset is 43 µm2. In Fig. 6, the mean apical surface area for the 48 hpf wild-type sibling OC dataset is 147 µm2, the mean apical surface area for the tbx5a mutant OC dataset is 108 µm2, and the mean Z-score for this experimental group is -0.67 (i.e. 0.67 standard deviations below the control mean). Therefore, for the apical surface area of tbx5a mutant OC cardiomyocytes,

mean norm.exp = 142 µm2 + (43 µm2 * -0.67) = 113 µm2

mean values were calculated in this way for apical surface area and apicobasal length for each dataset (myh6 OC, myh6 IC, tbx5a OC, and tbx5a IC) and plotted alongside the raw mean apical surface area and apicobasal length for the 37 hpf and 48 hpf wild-type OC and IC datasets from Fig. 1.

Statistical analysis, plotting, and figure design

Statistics were performed in R (v4.4.1) 101 and RStudio )v2025.09.1+401) 102. For all comparisons, statistical significance was determined by the Wilcoxon test, which does not assume normal data distribution. Statistically significant p-values are indicated in graphs by asterisks as follows: * denotes p<0.05; ** denotes p<0.01; *** denotes p<0.001. The exact number of embryos and/or cells examined for each experiment is reported in the corresponding figure legend.

The following R packages were used to restructure, statistically analyze, and plot the quantitative data captured from micrographs: dplyr 103, ggbeeswarm 104, ggplot2 105, ggpmisc 106, ggpubr 107, ggtext 108, Lattice 109, plyr 110, Tidyverse 111, and ggnewscale 112. In Figs. 1 and 4-7 and Supplementary Figs. 5-7 and 9, SuperPlots 113 were employed to portray variation between samples within the same experimental group. Figures were produced in Adobe InDesign v.18.5.2.107 and Inkscape v1.3.2.

Data availability

The data generated in this study are provided in the Source Data file.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In this study, Leeberg et al investigate the influence of the actomyosin network on cardiomyocyte cell shape during curvature morphogenesis, with specific emphasis on apicobasal organisation of the cytoskeleton. Using cardiomyocyte morphometric analysis at different times in cardiac development, the authors describe that OC and IC cells diverge in their morphology and show F-actin and p-Myosin organisation precedes cell shape changes, presumably driving ventricular wall curvature. Whilst this is perhaps unsurprising, it is novel and has (to our knowledge) not been demonstrated in this process before. Using pharmacological treatments, transgenics, and mutant analyses combined with cell transplant assays, they investigate the actomyosin network during this process and how it is regulated by both intrinsic and extrinsic factors. The experiments are elegantly performed and analyses carefully conducted, with the interpretation of the data mostly appropriate.\nThere are two major concerns with the work that require addressing:\n1. The work using the myl9 transgenics (DN and CA) are not convincing and at times contradictory. There are several aspects to this:\na. The generation of stable DN and CA myl9 transgenics is surprising. If these are major modifiers of the actomyosin network, it’s surprising they are viable (for e.g., sustained blebbistatin exposure would likely be lethal). Is there a phenotype in the adult fish? Are they healthy and 100% viable? Is there a difference in the curvature of the heart? If not, it could be argued that these are not the correct model to address central questions of the manuscript.\nb. DN-myl9 and CA-myl9 are used to investigate the role of NMII in OC and IC cardiomyocyte morphologies. DN-myl9 weakens the actomyosin machinery while CA-myl9 strengthens it. Yet, the use of both transgenes results in similar phenotypes such as decreased apical surface area and F-actin levels. Could the authors comment on this?\nc. What’s the explanation for the reduced fluorescence of F-actin? Could it be squelching or a similar effect that is different from intended?\nd. What are the expression levels of the transgenics like? Are they weaker than the mosaic or injected F0? Could there be some silencing/inhibition of the transgene occurring that might account for the subtlety of the phenotypes?\ne. Some effects in the mosaic analysis are opposite to that observed in the transgenic. For e.g. the circularity of stable CA myl9 is decreased in the OC and yet increased in the mosaic analysis. Could the authors comment on this?\n2. Whilst this is an elegant study, it is brief, which limits the contribution. Could the authors take the work a little further? For example, is there any information about what Tbx5a might be regulating to control actomyosin organisation? Are there any Formins, Rho GTPases, etc that are known to be downstream of Tbx5? Alternatively, the observation of the basal projections is intriguing and there is a lot of speculation in the Discussion about how this might be regulated. Could this be tested to some extent?\nMinor comments:\n- The authors show that there are cell shape differences between the OC and the IC. This appears to be driven by the organisation of the actomyosin network. While manipulation of the actomyosin network (by DN-myl9, CA-myl9, blood flow, tbx5a mutants etc.) results in reduced planar spread in OC cells, localisation of F-actin and in some cases pMyo in the proximal OC is the most affected. Looking more closely in WT hearts, the expression pattern of F-actin and pMyo (fig S3 and figS4D) in the distal OC appears more similar to the IC. This begs the question – are there cell shape differences between the proximal and distal cells of the OC? Ie. is the proximal OC more squamous than the distal OC? Additionally, in treated samples are proximal OC cell shape more disrupted?\n- The authors posit that changes in cell shape accompany curvature formation. In several assays conducted by the authors cell shape especially in the OC is affected but its effect on characteristic chamber curvatures is not measured. Could this be addressed?\n- Disruption of NMII, blood flow and tbx5a function result in reduced F-actin levels. Is this due to a reduction of actin expression or is actin polymerisation affected? An IF for G-actin or a western blot for actin may aid in answering this.\n- Apicobasal length is measured using confocal images. In some instances, it appears as though the images have been acquired with in the X-Y plane. At other times, apicobasal axis appears to be acquired in the Z-axis. Given that this axis can be distorted during acquisition, can the authors provide some assurance of the measurements? For example, can the authors confirm that for the measurements there is representation of the apicobasal plane imaged in either the X or Y and, importantly, that they are similar/consistent with those measured on the Z-plane?\n- What happens to mb upon blebbistatin treatment or in tbx5a mutants?\n- 3D reconstructions would aid in the visualisation of the varying cell shapes of the OC and IC.\n- Fig. S3E, F labelled as 50hpf (also in the figure legend) but is stated as 48hpf in text (pg. 9, line 1).\n- Page 14, last line “normal” should read “wildtype”", "reviewer_2": "In this manuscript, Leerberg et al. describe the cellular and actomyosin changes that occur in zebrafish ventricular cardiomyocytes between 36-48 hpf leading to divergence of the outer and inner curvatures (OC, IC). While there is already a substantial literature on this topic, what makes this study stand out is the high level of resolution and quantification, taking advantage of a host of valuable transgenic reporter, transgenic driver, and mutant strains. After defining the OC and IC domains based on nppa and mb expression, respectively, they show that from 36-48 hpf the OC cells expand along the planar axis, increasing apical surface area, while the IC cells expand mostly along the apicobasal axis, becoming taller and rounder. This is correlated primarily with F-actin enrichment at the basal surface of OC especially proximal near the AVC, while F-actin remains distributed throughout the IC cells. OC planar expansion is shown to be MNII dependent, and disrupted by dysregulation of myl9, lack of blood flow, or mutation of tbx5a. The study makes use of blastomere transplantation to show that at least some aspects of regulating actomyosin distribution are cell autonomous. Overall, this is an excellent description for how localized cellular distribution of the actomyosin machinery drives cellular morphology of OC cells leading to organ curvature. The authors also make some interesting observations on cellular protrusions that suggest a possible link to underlying ECM, although this is quite speculative.\nOverall, this is a comprehensive and highly detailed analysis of the early morphological events leading to heart looping at a critical developmental stage. It is entirely convincing and suitable for publication in Nature Communications. I have no major issues. The numbers of samples analyzed, and level of resolution is truly impressive. There are some caveats, since for imaging purposes the hearts are manipulated ex vivo, which probably should be mentioned clearly up front. It would also be helpful to indicate what eventually happens to these hearts (assuming they were not dissected but embryos left to develop). For example, presumably the DN-myl9 and CA-myl9 hearts go on to loop etc. since these appear to be established lines. Why do they recover? Likewise for the drug-treated larvae, do they recover? The authors should also check to align the text and figure panels for congruency (36, 37, 48, 50 hr?). These are very minor issues and the next most interesting mechanistic studies, such as identifying the tbx5-regulated genes that mediate these phenomena are arguably beyond the scope of this excellent paper.", "reviewer_3": "The study provides a comprehensive exploration of how actomyosin dynamics during cell shape changes in cardiomyocytes during ventricular chamber curvature formation. The findings that OC and IC cardiomyocytes display distinct actomyosin organization and morphologies—OC cells becoming squamous and expanding along the planar axis, while IC cells become cuboidal and elongate along the apicobasal axis—are insightful. These shape changes are preceded by specific actomyosin patterns and are influenced by both intrinsic factors (such as actomyosin and Tbx5a) and extrinsic cues (like blood flow). The methods section is thorough, and the complexity of the data adds to the study's depth. However, there are some aspects where the data remain inconclusive or unclear, particularly in the subcellular localization patterns of actomyosin due to limitations in image resolution and analysis. Since this paper addresses regional patterning, higher-quality images would improve the clarity of the findings.\nA significant concern I have is the lack of mention of the extracellular matrix (ECM), particularly cardiac jelly, as an extrinsic source of curvature. While the study focuses comprehensively on the intrinsic role of actomyosin, and references other extrinsic sources like blood flow, it would be beneficial to at least mention the ECM, as it likely plays a role in curvature formation, similar to its role in gut looping or atrioventricular canal (AVC) formation during the same developmental stages. Additionally, while actomyosin regional patterning may precede cell shape changes, these changes could still be a response to extrinsic forces (like the ECM) rather than being causal to curvature induction. This distinction should be addressed more explicitly to clarify the contribution of actomyosin dynamics.\nThe mechanism underlying the differences in morphology between OC and IC cells in response to changes in actomyosin activity also remains unclear. The expected roles of dominant-negative and constitutively active myosin variants, disrupting or increasing contractility, need further clarification, especially in terms of how these changes lead to the observed elongation and cuboidal phenotypes.\nI recommend that the number of replicates for each experiment be stated clearly.\nSpecific Suggestions:\nFigure 1:\n• Page 8, line 6: The statement \"Cells allocate their 'new volume' to different axes\" raises the question of whether cell division is occurring. If it is, it would be helpful to address whether cell division rates are comparable in OC and IC cells.\n• Page 9, lines 30–31; Page 10, line 1: It would be valuable to include a side-by-side comparison of actomyosin localization and cell shape divergence over time within the same embryos and cells. This would help to better connect the spatial and temporal patterns of actomyosin dynamics with the observed morphological changes.\nFigure S6:\nThe authors suggest that actomyosin is involved in planar expansion, but traditionally, higher actomyosin levels are associated with contraction, especially when non-muscle myosin II (NMII) is involved (as mentioned in the discussion). The result showing that Bleb treatment causes excessive planar expansion could be explained by the myosin motor being locked to F-actin, behaving more like a cross-linker rather than a motor. This suggests that myosin may be playing a contractile role rather than an expansive one.\nFigure 3:\nThe data presented here are not conclusive, and the differences between the experimental conditions seem too small, especially given that the quantification was done manually. It would be helpful to express the mutant transgenes in conditions where Rho kinase (Rok) or MLCK (or another kinase involved here) is knocked down, as the DN and CA versions of myosin may not dilute endogenous wild-type myosin sufficiently.\nFigure S7:\nThe effect on F-actin is quite surprising. The justification for the quantitative comparison should be more robust—were the immunostaining experiments performed on samples processed in the same way? Are the transgenes in the same genetic background? It would be useful to use a Utr(ABD) line or another method to confirm the findings\nFigure 4:\nThe authors claim that transplantation addresses the cell-intrinsic versus tissue-scale role of actomyosin; however, the transplantation leads to large groups of mutant cells rather than the salt-pepper labeling that would allow for a more precise evaluation. Using plasmid injection (rather than transplantation) may be more revealing of whether neighboring wild-type cells influence the shape of the mutant cells or not.\nFigure 5:\nThe findings regarding the total levels of F-actin remain unclear. A more detailed explanation and justification for the quantitative comparison would be helpful.\nFigure 8:\nIncluding an illustration or phase diagram that summarizes the effects of various actomyosin and Tbx perturbations on apical-basal length and apical surface area would greatly enhance the clarity and the working model."}, "author_response": "Response to Reviewers' Comments – Leerberg et al., NCOMMS-25-33229\nWe are very grateful to all three of the reviewers for their positive feedback regarding our manuscript.\nWe also greatly appreciate the reviewers' thoughtful suggestions for strategies to strengthen our\nmanuscript's value. We have now modified our submission in accordance with their input, both by\nadding new data and by adjusting the manuscript. Our revised submission includes five new\nsupplementary figures (Supplementary Figures 8, 10, 11, 12, 13), as well as adapted versions of previous\nfigures (Figures 3-7 and Supplementary Figures 7 and 9). We have highlighted the new and updated\ncontent in our revised manuscript and figure legends, although we have not highlighted formatting\nchanges (such as renumbered figures or relabeled panels). Altogether, we feel that the suggested changes\nhave substantially enhanced the significance and clarity of our manuscript, and we thank the reviewers\nfor their assistance with these improvements. Our point-by-point responses to the reviewers' comments\nare assembled below, with the reviewer comments represented verbatim in blue text and our responses\nprovided in black text.\nReviewer #1 (Remarks to the Author):\nIn this study, Leeberg et al investigate the influence of the actomyosin network on cardiomyocyte cell\nshape during curvature morphogenesis, with specific emphasis on apicobasal organisation of the\ncytoskeleton. Using cardiomyocyte morphometric analysis at different times in cardiac development, the\nauthors describe that OC and IC cells diverge in their morphology and show F-actin and p-Myosin\norganisation precedes cell shape changes, presumably driving ventricular wall curvature. Whilst this is\nperhaps unsurprising, it is novel and has (to our knowledge) not been demonstrated in this process\nbefore. Using pharmacological treatments, transgenics, and mutant analyses combined with cell\ntransplant assays, they investigate the actomyosin network during this process and how it is regulated by\nboth intrinsic and extrinsic factors. The experiments are elegantly performed and analyses carefully\nconducted, with the interpretation of the data mostly appropriate.\nWe are grateful to Reviewer #1 for their positive assessment of the novelty of our work and the quality\nof our experimental design, execution, and analysis.\nThere are two major concerns with the work that require addressing:\n1. The work using the myl9 transgenics (DN and CA) are not convincing and at times contradictory.\nThere are several aspects to this:\nWe thank the reviewer for their series of important questions regarding our use of the DN-myl9 and CA-\nmyl9 transgenes. As noted below, Reviewers #2 and #3 also asked questions about these transgenes, and\nwe greatly appreciate the opportunity to clarify the utility of these tools for our studies.\na. The generation of stable DN and CA myl9 transgenics is surprising. If these are major modifiers of\nthe actomyosin network, it’s surprising they are viable (for e.g., sustained blebbistatin exposure would\nlikely be lethal). Is there a phenotype in the adult fish? Are they healthy and 100% viable? Is there a\ndifference in the curvature of the heart? If not, it could be argued that these are not the correct model to\naddress central questions of the manuscript.\nThe stable transgenic lines Tg(myl7:DN-myl9-eGFP)bns333Tg and Tg(myl7:CA-myl9-eGFP)bns332Tg were\ngenerated by our co-authors Drs. Priya and Stainier and have been utilized in their previous publications\n(Priya et al., Nature, 2020; Albu et al., Nature Communications, 2024). Although cardiac chamber\nmorphology and function have not been carefully examined in adult transgenic fish, these prior studies\nhave established that adult transgenics are generally healthy, viable, and able to reproduce. It is\nimportant to note that these transgenes are expressed only in the myocardium, and so they do not affect\nthe cytoskeletal machinery in other tissues, as treatment with Blebbistatin would. Even so, we agree that\nit would be surprising for transgenics expressing very high levels of myocardial DN-myl9 or CA-myl9\nexpression to be viable. We therefore presume that the impact on the myocardial actomyosin network in\nthe Tg(myl7:DN-myl9-eGFP)bns333Tg and Tg(myl7:CA-myl9-eGFP)bns332Tg transgenic lines is relatively\nmodest. Indeed, our revised manuscript includes new data indicating that, while pMyosin levels are\nreduced in DN-myl9 hearts, some pMyosin still remains (Supplementary Fig. 8; p. 11); perhaps the\nsurvival of DN-myl9 fish is due to this residual non-muscle myosin II (NMII) activity. Despite the\npresence of residual NMII activity in DN-myl9 transgenic hearts, we think that the effect of DN-myl9\nexpression on both the levels and organization of F-actin (Supplementary Figs. 8 and 9; p. 11) position\nthe Tg(myl7:DN-myl9-eGFP)bns333Tg transgenic line as a reasonable and valuable tool for investigating\nthe influence of the actomyosin cytoskeleton on ventricular cardiomyocyte morphology. (As described\nin the next response below, we have decided to remove experiments utilizing Tg(myl7:CA-myl9-\neGFP)bns332Tg from this manuscript.)\nb. DN-myl9 and CA-myl9 are used to investigate the role of NMII in OC and IC cardiomyocyte\nmorphologies. DN-myl9 weakens the actomyosin machinery while CA-myl9 strengthens it. Yet, the use\nof both transgenes results in similar phenotypes such as decreased apical surface area and F-actin levels.\nCould the authors comment on this?\nThis is an interesting question, and its answer probably relates to the multiple cellular mechanisms at\nplay. For example, cardiomyocyte morphologies are likely to be influenced both by the cell-intrinsic\nroles of the actomyosin machinery and by the cell-extrinsic forces exerted upon a cell by its neighbors.\nIn transgenic hearts, both the intrinsic and extrinsic mechanisms are altered, and the differential\ninterplay between these components could yield decreased apical surface area of OC cardiomyocytes in\nboth the DN-myl9 and CA-myl9 contexts. Additionally, as shown in Supplementary Fig. 6, the timing of\nactomyosin modulation is an important factor, with NMII activity seeming to play different roles\nbetween 24-36 hpf and between 36-48 hpf. Since the myl7 promoter drives transgene expression\nthroughout the stages examined, it is possible that the effects of transgene expression during the later\ninterval could mask detection of earlier effects (or vice versa) (p. 21).\nTaking into account this input from Reviewer #1, together with related feedback from Reviewer #3\n(discussed below), we have decided to remove all experiments utilizing the CA-myl9 transgene from our\nmanuscript in order to streamline its overall narrative. Although we feel that the CA-myl9 data highlight\ninteresting aspects of the relevant cell biology, we recognize that their interpretation is complex and that\nthey are not needed to support our primary conclusions. In our revised manuscript (Fig. 3 and\nSupplementary Figs. 7-10; pp. 10-12), we focus on the use of the DN-myl9 transgene to demonstrate that\nreduction of NMII activity alters F-actin organization and inhibits the planar expansion of OC\ncardiomyocytes.\nc. What’s the explanation for the reduced fluorescence of F-actin? Could it be squelching or a similar\neffect that is different from intended?\nWe appreciate Reviewer #1's curiosity about the reduced phalloidin signal observed in hearts expressing\nDN-myl9; this is echoed below in Reviewer #1's third minor comment, as well as in two comments from\nReviewer #3. Although our studies have not yet elucidated the precise mechanisms responsible for the\nreduced levels of phalloidin signal in DN-myl9 transgenic hearts, we do not think that this observation is\nan imaging artifact. All hearts were treated and imaged similarly, regardless of experimental group, and\ncare was taken to avoid loss of fluorescent signal (e.g. avoiding unnecessarily long or intense laser\nexposure). Additionally, our revised manuscript includes new data that help to address this topic\n(Supplementary Fig. 8; p. 11). Specifically, in addition to exhibiting reduced levels of F-actin, DN-myl9\ntransgenic hearts display a coincident reduction in pMyosin levels (suggesting reduced NMII activity)\nand, by contrast, display levels of G-actin that are comparable to the levels in wild-type siblings\n(Supplementary Fig. 8). These new data suggest that, although actin monomers are readily available in\nDN-myl9 transgenic hearts, they are not being polymerized into filaments to the same degree as in wild-\ntype siblings (p. 11). This is possibly due to the reduced NMII activity in DN-myl9 hearts, as NMII is\nknown to affect the F-actin landscape in myriad ways (e.g. references #68-#76 in our revised\nmanuscript).\nd. What are the expression levels of the transgenics like? Are they weaker than the mosaic or injected\nF0? Could there be some silencing/inhibition of the transgene occurring that might account for the\nsubtlety of the phenotypes?\nIn the transgenic line Tg(myl7:DN-myl9-eGFP)bns333Tg, the DN-myl9 protein is fused to eGFP, and we\nfind that eGFP fluorescence is easily visible in the heart by ~24 hpf and that fluorescence levels are\nreasonably consistent among transgenic embryos. Additionally, this level of eGFP fluorescence appears\ncomparable to the level seen in donor-derived transgenic cardiomyocytes in mosaic hearts generated via\nblastomere transplantation (Fig. 3). In contrast, transgene injection generates F0 embryos with mosaic\nhearts that exhibit highly variable amounts of fluorescence from cell to cell (Supplementary Fig. 10);\nthis broad range of mosaicism is common in F0 zebrafish embryos (e.g. Fisher et al., Nature Protocols,\n2006). Therefore, we agree with Reviewer #1 that the level of DN-myl9 expression in our transgenic\nembryos is lower than can sometimes be found in injected F0 embryos. As discussed above (Reviewer\n#1, comment #1a), we suspect that these relatively modest DN-myl9 expression levels are likely to\naccount for the viability of Tg(myl7:DN-myl9-eGFP)bns333Tg transgenic embryos. Although higher levels\nof expression can be achieved in injected F0 embryos, we feel that the consistency of DN-myl9\nexpression levels found in transgenic embryos is valuable for our studies. Even so, motivated by the\ninput from Reviewer #1 and Reviewer #3, our revised manuscript also incorporates new data in which\nwe complement our blastomere transplantation approach with analysis of transgene-injected F0 embryos\n(Supplementary Fig. 10; p. 12). Satisfyingly, we found that transgene-expressing OC cardiomyocytes in\ninjected F0 embryos exhibited planar expansion defects similar to the defects found in the donor-derived\nOC cardiomyocytes in our blastomere transplantation experiments (Fig. 3 and Supplementary Fig. 10; p.\n12).\ne. Some effects in the mosaic analysis are opposite to that observed in the transgenic. For e.g. the\ncircularity of stable CA myl9 is decreased in the OC and yet increased in the mosaic analysis. Could the\nauthors comment on this?\nAs alluded to above (Reviewer #1, comment #1b), differences between cell morphologies in transgenic\nhearts and cell morphologies in mosaic hearts could reflect the differential impacts of the cell-intrinsic\nand cell-extrinsic roles played by the actomyosin machinery. In a transgenic heart, all neighboring\ncardiomyocytes express the transgene, providing less (DN-myl9) or more (CA-myl9) NMII activity than\nwild-type neighbors would. Even so, we note that the effects of DN-myl9 expression are relatively\ncomparable in transgenic hearts (Supplementary Fig. 7) and in mosaic hearts (Fig. 3): in both scenarios,\ninhibition of NMII activity reduces the planar expansion of OC cardiomyocytes.\nSince the combination of cell-intrinsic and cell-extrinsic influences complicates the interpretation of\nphenotypes seen in transgenic hearts, we have decided to emphasize the significance of our DN-myl9\nmosaic analysis (Fig. 3) in our revised manuscript, and we have moved our analysis of DN-myl9\ntransgenic hearts to the Supplementary Material (Supplementary Fig. 7). Additionally, as mentioned\nabove (Reviewer #1, comment #1b), we have decided to remove all experiments utilizing the CA-myl9\ntransgene from our manuscript in order to streamline its overall narrative.\n2. Whilst this is an elegant study, it is brief, which limits the contribution. Could the authors take the\nwork a little further? For example, is there any information about what Tbx5a might be regulating to\ncontrol actomyosin organisation? Are there any Formins, Rho GTPases, etc that are known to be\ndownstream of Tbx5? Alternatively, the observation of the basal projections is intriguing and there is a\nlot of speculation in the Discussion about how this might be regulated. Could this be tested to some\nextent?\nWe greatly appreciate Reviewer #1’s interest in these important future directions for our studies. We are\ncertainly keen to understand the factors that act downstream of Tbx5a to control actomyosin\norganization, and we note that existing transcriptomic analyses do suggest that certain actomyosin-\nassociated genes, such as EZRIN (Kathiriya et al., Developmental Cell, 2021) or formin like 2b (Pawlak\net al., Genome Research, 2019), could be regulated by Tbx5. Although we are actively investigating\npotential downstream factors, thorough analysis of each candidate will require a significant amount of\nadditional experimentation, beyond the scope and timeframe for revision of our current manuscript.\nWe therefore respectfully decline to include this research direction within our revised submission. Similarly,\nwe appreciate Reviewer #1’s curiosity about the regulation of the basal projections shown in\nSupplementary Figures 16 and 17. We are actively investigating this open question, and we believe that\nit may ultimately relate to the questions raised by Reviewer #3 regarding ECM involvement (see below).\nHowever, the mechanisms underlying these intriguing cell behaviors are likely to be complex, and their\nelucidation will require extensive experimentation that lies beyond the scope of this revision. Again, we\nrespectfully decline to include this research direction in our revised manuscript.\nMinor comments:\n- The authors show that there are cell shape differences between the OC and the IC. This appears to be\ndriven by the organisation of the actomyosin network. While manipulation of the actomyosin network\n(by DN-myl9, CA-myl9, blood flow, tbx5a mutants etc.) results in reduced planar spread in OC cells,\nlocalisation of F-actin and in some cases pMyo in the proximal OC is the most affected. Looking more\nclosely in WT hearts, the expression pattern of F-actin and pMyo (fig S3 and figS4D) in the distal OC\nappears more similar to the IC. This begs the question – are there cell shape differences between the\nproximal and distal cells of the OC? Ie. is the proximal OC more squamous than the distal OC?\nAdditionally, in treated samples are proximal OC cell shape more disrupted?\nWe thank Reviewer #1 for asking about this interesting topic. We have not detected any significant and\nconsistent morphometric differences between wild-type OC cardiomyocytes in the regions that we\ndefine as proximal and distal at 48 hpf, and we mention this in our revised Discussion (p. 22). However,\nit is important to note that the cardiomyocytes found in the regions that we define as proximal and distal\nwhen analyzing cell morphologies at 48 hpf may not precisely correspond to the cardiomyocytes found\nin the regions that we define as proximal and distal when analyzing actomyosin organization at 36 hpf,\nwhich may mask our detection of meaningful morphological differences at 48 hpf. In future studies\nbeyond the scope of the current manuscript, we plan to utilize timelapse imaging to track these\npopulations of cells between 36 and 48 hpf.\nAlthough we have not found significant differences in the morphologies of proximal and distal OC\ncardiomyocytes in wild-type hearts, we have also analyzed whether proximal and distal OC cells are\ndifferentially impacted in different genetic backgrounds. We did not observe significant morphometric\ndifferences between proximal and distal OC cells in DN-myl9 transgenic or tbx5a mutant hearts (data\nnot shown). However, in myh6 mutant hearts, proximal OC cardiomyocytes did seem to be more\nseverely affected than distal OC cardiomyocytes (Reviewer Fig. 1). While apical surface area is affected\nsimilarly in proximal and distal regions (Reviewer Fig. 1a; Cohen’s d = 0.61 (proximal) vs 0.72 (distal),\nD 0.11), apicobasal length is more affected in the proximal region (Reviewer Fig. 1b; Cohen’s d = -0.55\n(proximal) vs -0.27 (distal), D 0.28), as is apical circularity (Reviewer Fig. 1c; Cohen’s d = 0.53\n(proximal) vs -0.05 (distal), D 0.58). This analysis suggests that the proximal and distal regions of the\nOC might be differentially sensitive to the biomechanical cues associated with blood flow, perhaps due\nto underlying differences in the actomyosin landscape. While we are happy to share this information\nwith the reviewers, we prefer not to incorporate it into our revised manuscript at this time, since we feel\nthat substantially more work will be required to understand why we only see these differences in the\nmyh6 mutant context (and not in the tbx5a mutant or DN-myl9 transgenic contexts) and to understand\nwhy the differential effects are found for apicobasal length and circularity (and not for apical surface\narea).\nReviewer Figure 1. Blood flow affects cardiomyocytes in the proximal and distal regions of the OC\ndifferently.\n(a-c) OC data from Fig. 4m-o, split into proximal and distal regions. For all violin plots, each dot\nrepresents an individual cell, and each black bar represents the mean. * denotes p < 0.05 and ***\ndenotes p < 0.001, Wilcoxon test. Cohen’s d values represent the effect size. Wild-type proximal OC\n(N=5 embryos, 142 cells); myh6 proximal OC (N=6 embryos, n=147 cells); wild-type distal OC (N=5\nembryos, 125 cells); myh6 distal OC (N=6 embryos, n=128 cells).\n- The authors posit that changes in cell shape accompany curvature formation. In several assays\nconducted by the authors cell shape especially in the OC is affected but its effect on characteristic\nchamber curvatures is not measured. Could this be addressed?\nWe agree with Reviewer #1 that it would be desirable to evaluate how individual cardiomyocyte\nmorphologies contribute to the overall geometry of the ventricle, and we plan to address this in future\nstudies. This work will involve the creation of new two-dimensional and three-dimensional metrics for\nchamber curvatures, establishment of the variation found in wild-type curvatures at stages between 24\nand 48 hpf, and mathematical modeling of the contributions of cell shapes to curvature dimensions;\ntherefore, this combination of pursuits lies beyond the scope of our current manuscript.\n- Disruption of NMII, blood flow and tbx5a function result in reduced F-actin levels. Is this due to a\nreduction of actin expression or is actin polymerisation affected? An IF for G-actin or a western blot for\nactin may aid in answering this.\nWe thank Reviewer #1 (and Reviewer #3, below) for encouraging us to investigate this topic further. We\nappreciated and pursued the suggestion to examine G-actin levels, and this has led to the inclusion of\nthree new figures in our revised manuscript (Supplementary Figs. 8, 12, and 13). As discussed above\n(Reviewer #1, major comment #1c), we found that the reduced levels of F-actin in DN-myl9 transgenic\nhearts are accompanied by levels of G-actin that are comparable to the levels in wild-type siblings\n(Supplementary Fig. 8). These data suggest that, although actin monomers are readily available in DN-\nmyl9 transgenic hearts, they are not being polymerized into filaments to the same degree as in wild-type\nsiblings (p. 11). Similarly, we have found that myh6 mutants and tbx5a mutants exhibit reduced levels of\nF-actin, together with G-actin levels that are similar to wild-type (Supplementary Figs. 12 and 13). As in\nDN-myl9 transgenics, these data suggest that availability of actin monomers is normal in myh6 mutants\nand in tbx5a mutants, but that actin modifiers are not leading to normal polymerization of actin\nfilaments in these hearts (pp. 13, 14, 20).\n- Apicobasal length is measured using confocal images. In some instances, it appears as though the\nimages have been acquired with in the X-Y plane. At other times, apicobasal axis appears to be acquired\nin the Z-axis. Given that this axis can be distorted during acquisition, can the authors provide some\nassurance of the measurements? For example, can the authors confirm that for the measurements there is\nrepresentation of the apicobasal plane imaged in either the X or Y and, importantly, that they are\nsimilar/consistent with those measured on the Z-plane?\nWe thank Reviewer #1 for raising this important point. Our standard practice is to mount hearts laterally,\nwith one side of the ventricle flat against the coverslip, and we have clarified this in our revised Methods\nsection (p. 26). In this orientation (Reviewer Fig. 2a,b), the X-Y axes of most of the OC and IC cells are\nperpendicular to the objective. This results in more resolvable signal in the section views used to\nmeasure apicobasal length (Reviewer Fig. 2b), compared to when hearts are mounted with the OC\nclosest to the objective (Reviewer Fig. 2c,d). Additionally, mounting in our standard orientation allows\nus to visualize both the OC and IC with similar clarity and intensity (compare the ICs in Reviewer Fig.\n2b and 2d). However, it is important to note that the cells in the curvatures are not all in the same plane,\nas the ventricle is a bulging, three-dimensional structure. Therefore, we performed an experiment in\nwhich the same three hearts were imaged in two perpendicular orientations, and we measured the\napicobasal lengths of the OC cells along the midline in each set of images (Reviewer Fig. 2e).\nReassuringly, we found similar apicobasal length values when comparing these perpendicular\norientations (Reviewer Fig. 2e).\nReviewer Figure 2. Hearts imaged in perpendicular orientations reveal similar apicobasal lengths\nof OC cardiomyocytes.\n(a,b) 3D reconstruction of the OC (a) and section through the ventricle (b) of a heart mounted flat\nagainst the coverslip, resulting in one side of the ventricle being closest to the objective. (c,d) 3D\nreconstruction of the OC (c) and section through the ventricle (d) of the same heart shown in (a) and (b),\nbut mounted with the OC against the coverslip and closest to the objective. Sections in (b) and (d) are\ntaken at the positions represented by the dotted red lines in (a) and (c). Scale bars = 20 μm. (e) Three\nhearts were imaged in two orientations each. For each of the three hearts, apicobasal (AB) length was\nmeasured (as in Fig. 1) for OC cardiomyocytes that intersected with the midline (represented by the\ndotted red line in (a) and (c)). In the violin plots, each heart is represented by a different shape; and each\ncell measured is represented by a different color. p > 0.05, paired Wilcoxon test.\n- What happens to mb upon blebbistatin treatment or in tbx5a mutants?\nWe have not noted any differences in the pattern of mb (myoglobin) expression in the tbx5a mutant\nventricle at 36 hpf, suggesting that the initial patterning of IC identity may not depend upon tbx5a\nfunction. We have not examined mb expression in blebbistatin-treated embryos, and we chose not to\nprioritize this experiment as part of our revision efforts for this manuscript.\n- 3D reconstructions would aid in the visualisation of the varying cell shapes of the OC and IC.\nWe appreciate Reviewer #1’s interest in viewing 3D reconstructions of the morphologies of individual\nOC and IC cardiomyocytes. However, the methods used in the relevant experiments (Figs. 1, 4, 6) do\nnot permit this type of visualization: these studies used Cdh2 to label the lateral membranes of\ncardiomyocytes but did not include a transgene like Tg(myl7:eGFP-Hsa.HRAS) that outlines all of the\ncardiomyocyte surfaces. We have therefore respectfully declined to include 3D cardiomyocyte\nmorphology reconstructions in our revised manuscript.\n- Fig. S3E, F labelled as 50hpf (also in the figure legend) but is stated as 48hpf in text (pg. 9, line 1).\nWe have made this correction in our revised manuscript (p. 8). Please note that the panel labeling for\nSupplementary Fig. 3 has changed, but we have changed \"48 hpf\" to \"50 hpf\" as needed.\n- Page 14, last line “normal” should read “wildtype”\nWe have made this correction in our revised manuscript (p. 14).\nReviewer #2 (Remarks to the Author):\nIn this manuscript, Leerberg et al. describe the cellular and actomyosin changes that occur in zebrafish\nventricular cardiomyocytes between 36-48 hpf leading to divergence of the outer and inner curvatures\n(OC, IC). While there is already a substantial literature on this topic, what makes this study stand out is\nthe high level of resolution and quantification, taking advantage of a host of valuable transgenic\nreporter, transgenic driver, and mutant strains. After defining the OC and IC domains based on nppa and\nmb expression, respectively, they show that from 36-48 hpf the OC cells expand along the planar axis,\nincreasing apical surface area, while the IC cells expand mostly along the apicobasal axis, becoming\ntaller and rounder. This is correlated primarily with F-actin enrichment at the basal surface of OC\nespecially proximal near the AVC, while F-actin remains distributed throughout the IC cells. OC planar\nexpansion is shown to be MNII dependent, and disrupted by dysregulation of myl9, lack of blood flow,\nor mutation of tbx5a. The study makes use of blastomere transplantation to show that at least some\naspects of regulating actomyosin distribution are cell autonomous. Overall, this is an excellent\ndescription for how localized cellular distribution of the actomyosin machinery drives cellular\nmorphology of OC cells leading to organ curvature. The authors also make some interesting\nobservations on cellular protrusions that suggest a possible link to underlying ECM, although this is\nquite speculative.\nOverall, this is a comprehensive and highly detailed analysis of the early morphological events leading\nto heart looping at a critical developmental stage. It is entirely convincing and suitable for publication in\nNature Communications. I have no major issues. The numbers of samples analyzed, and level of\nresolution is truly impressive. There are some caveats, since for imaging purposes the hearts are\nmanipulated ex vivo, which probably should be mentioned clearly up front. It would also be helpful to\nindicate what eventually happens to these hearts (assuming they were not dissected but embryos left to\ndevelop). For example, presumably the DN-myl9 and CA-myl9 hearts go on to loop etc. since these\nappear to be established lines. Why do they recover? Likewise for the drug-treated larvae, do they\nrecover? The authors should also check to align the text and figure panels for congruency (36, 37, 48, 50\nhr?). These are very minor issues and the next most interesting mechanistic studies, such as identifying\nthe tbx5-regulated genes that mediate these phenomena are arguably beyond the scope of this excellent\npaper.\nWe are grateful to Reviewer #2 for their strong support of the quality, rigor, and significance of our\nstudies. We also appreciate the minor issues raised by Reviewer #2, and we can address each of these,\nas follows:\n- We have clarified that hearts are dissected from embryos before imaging, both in our Methods section\n(p. 26) and in the Fig. 1 legend.\n- Above (Reviewer #1, comment #1a), we have discussed that DN-myl9 transgenic hearts likely recover\ndue to their remaining pMyosin (Supplemental Fig. 8).\n- We have not performed washout experiments to examine how well Latrunculin B-treated or\nBlebbistatin-treated embryos recover. However, since these drugs affect actin or myosin dynamics\nthroughout the embryo, it is likely that their recovery could be hindered by defects in multiple tissues.\n- We have corrected issues in the congruency between our text and figure panels.\nReviewer #3 (Remarks to the Author):\nThe study provides a comprehensive exploration of how actomyosin dynamics during cell shape\nchanges in cardiomyocytes during ventricular chamber curvature formation. The findings that OC and\nIC cardiomyocytes display distinct actomyosin organization and morphologies—OC cells becoming\nsquamous and expanding along the planar axis, while IC cells become cuboidal and elongate along the\napicobasal axis—are insightful. These shape changes are preceded by specific actomyosin patterns and\nare influenced by both intrinsic factors (such as actomyosin and Tbx5a) and extrinsic cues (like blood\nflow). The methods section is thorough, and the complexity of the data adds to the study's depth.\nHowever, there are some aspects where the data remain inconclusive or unclear, particularly in the\nsubcellular localization patterns of actomyosin due to limitations in image resolution and analysis. Since\nthis paper addresses regional patterning, higher-quality images would improve the clarity of the\nfindings.\nWe are grateful to Reviewer #3 for their positive evaluation of the “comprehensive”, “insightful”, and\n“thorough” nature of our studies, and we also appreciate their suggestions for ways to improve the\nclarity and impact of our data. Regarding Reviewer #3’s request for higher-quality images, we have\nadjusted the sizing of several figures in our revised manuscript (Figs. 1, 2, 3, 4, 6) in an effort to\nimprove the visibility of their details.\nA significant concern I have is the lack of mention of the extracellular matrix (ECM), particularly\ncardiac jelly, as an extrinsic source of curvature. While the study focuses comprehensively on the\nintrinsic role of actomyosin, and references other extrinsic sources like blood flow, it would be\nbeneficial to at least mention the ECM, as it likely plays a role in curvature formation, similar to its role\nin gut looping or atrioventricular canal (AVC) formation during the same developmental stages.\nWe appreciate Reviewer #3’s suggestion, and we agree that the ECM likely plays an important role\nduring curvature formation and the underlying cellular and subcellular mechanisms. In our revised\nmanuscript, we have added text to the Discussion to highlight this likelihood (pp. 18, 20, 22).\nAdditionally, while actomyosin regional patterning may precede cell shape changes, these changes could\nstill be a response to extrinsic forces (like the ECM) rather than being causal to curvature induction.\nThis distinction should be addressed more explicitly to clarify the contribution of actomyosin dynamics.\nWe agree with Reviewer #3 that extrinsic forces, including the ECM, influence the cytoskeleton, and it\nis certainly possible that the observed differences in cytoskeletal organization, although correlated with\nshape change, are not causative. In our revised manuscript, we have added text to the Discussion to\naddress these possibilities and the future work that will be needed to test them (pp. 18, 20, 22).\nThe mechanism underlying the differences in morphology between OC and IC cells in response to\nchanges in actomyosin activity also remains unclear. The expected roles of dominant-negative and\nconstitutively active myosin variants, disrupting or increasing contractility, need further clarification,\nespecially in terms of how these changes lead to the observed elongation and cuboidal phenotypes.\nPlease see responses above to Reviewer #1, comments #1a, #1b, and #1c. Notably, our revised\nmanuscript includes new data demonstrating that pMyosin levels are reduced in DN-myl9 hearts,\n(Supplemental Fig. 8). Also, as discussed above, we have decided to remove all experiments utilizing\nthe CA-myl9 transgene from our revised manuscript.\nI recommend that the number of replicates for each experiment be stated clearly.\nWe thank Reviewer #3 for this suggestion. In our revised manuscript, all relevant figure legends now\ninclude the number of experimental replicates, along with the number of embryos analyzed and, where\nappropriate, the number of cells analyzed.\nSpecific Suggestions:\nFigure 1:\n• Page 8, line 6: The statement \"Cells allocate their 'new volume' to different axes\" raises the question of\nwhether cell division is occurring. If it is, it would be helpful to address whether cell division rates are\ncomparable in OC and IC cells.\nWe did not evaluate cell division rates during this set of studies. However, prior work has shown that\nthere is very little proliferation in the zebrafish myocardium between 24-48 hpf (de Pater et al., 2009).\nSpecifically, this study found only <2 phospho-His+ cardiomyocytes per heart at 30, 36, and 48 hpf, and\nBrdU treatment between 24-48 hpf only yielded ~16 BrdU+ cardiomyocytes per heart, with more than\nhalf of these located near the cardiac poles. Therefore, it is unlikely that cell division has much of an\nimpact on OC and IC cells during the ~36-48 hpf timeframe that is the focus of our manuscript.\n• Page 9, lines 30–31; Page 10, line 1: It would be valuable to include a side-by-side comparison of\nactomyosin localization and cell shape divergence over time within the same embryos and cells. This\nwould help to better connect the spatial and temporal patterns of actomyosin dynamics with the\nobserved morphological changes.\nWe thank Reviewer #3 for this suggestion, and we certainly appreciate the value of being able to follow\nactomyosin organization and cell shape changes simultaneously in the same cells over time.\nUnfortunately, our current imaging tools and techniques cannot accommodate the live imaging\nexperiments necessary for these side-by-side comparisons, as the ventricle is located deep under the\nhead during the critical stages of actomyosin dynamics studied here. However, in future studies,\nbeyond the scope of our current manuscript, we hope to pursue this direction by adopting newly emerging\ntechniques, such as Dr. Priya’s “body explant” strategy (Jie et al., bioRxiv, 2025).\nFigure S6:\nThe authors suggest that actomyosin is involved in planar expansion, but traditionally, higher\nactomyosin levels are associated with contraction, especially when non-muscle myosin II (NMII) is\ninvolved (as mentioned in the discussion). The result showing that Bleb treatment causes excessive\nplanar expansion could be explained by the myosin motor being locked to F-actin, behaving more like a\ncross-linker rather than a motor. This suggests that myosin may be playing a contractile role rather than\nan expansive one.\nReviewer #3 brings up an interesting point regarding the different potential roles of NMII. We think that\nour data support multiple roles for NMII (pp. 10, 21): an earlier role (24-36 hpf) when NMII is involved\nin constructing the F-actin landscape and actualizing the subsequent divergence of OC and IC cell\nshapes, and a later role (36-48 hpf) when NMII helps maintain cell shape and size via its contractile\nproperties (as Reviewer #3 suggests). This idea is supported by the stage-dependent nature of our results\nwith Blebbistatin: treatment from 24-36 hpf resulted in smaller cells at 48 hpf, but treatment from 36-48\nhpf resulted in larger cells at 48 hpf (p. 10). It will be important in future work to be able to temporally\ncontrol cytoskeletal manipulations to better understand these complexities (p. 21).\nFigure 3:\nThe data presented here are not conclusive, and the differences between the experimental conditions\nseem too small, especially given that the quantification was done manually. It would be helpful to\nexpress the mutant transgenes in conditions where Rho kinase (Rok) or MLCK (or another kinase\ninvolved here) is knocked down, as the DN and CA versions of myosin may not dilute endogenous wild-\ntype myosin sufficiently.\nWe appreciate Reviewer #3’s interest in this topic, which is shared with Reviewer #1. We agree that the\nimpact of the DN-myl9 transgene on NMII activity is an important question. As discussed above in\nresponse to Reviewer #1, comment #1a, we have addressed this by examining the levels of pMyosin in\nDN-myl9 transgenic hearts, where we found a significant reduction in comparison to wild-type hearts\n(Supplementary Fig. 8; p. 11). We reason that this reduction, coupled with the observed effects of the\nDN-myl9 transgene on total F-actin and on F-actin organization (Supplementary Figs. 8 and 9; p. 11),\npositions the DN-myl9 transgene as a valuable tool for investigating the influence of the actomyosin\ncytoskeleton on ventricular cardiomyocyte morphology. Additionally, in our revised submission, we\nhave decided to emphasize the significance of our DN-myl9 mosaic analysis (now Fig. 3 of our revised\nmanuscript), and we have moved our analysis of DN-myl9 transgenic hearts to the Supplementary\nMaterial (Supplementary Fig. 7). Finally, as mentioned above (Reviewer #1, comment #1b), we have\ndecided to remove all experiments utilizing the CA-myl9 transgene from our manuscript in order to\nstreamline its overall narrative.\nFigure S7:\nThe effect on F-actin is quite surprising. The justification for the quantitative comparison should be\nmore robust—were the immunostaining experiments performed on samples processed in the same way?\nAre the transgenes in the same genetic background? It would be useful to use a Utr(ABD) line or\nanother method to confirm the findings\nWe appreciate Reviewer #3’s interest in this point, which is again shared with Reviewer #1 (comment\n#1c above). In our revised manuscript, we address the F-actin levels in DN-myl9 transgenics in\nSupplementary Fig. 8. The samples examined in Supplementary Fig. 8 were indeed processed in a\nparallel fashion (see Methods description on pp. 23-25), and the DN-myl9 transgenic embryos and their\nwild-type siblings were in the same genetic background. As described above (Reviewer #1, comment\n#1c), we found that in addition to exhibiting reduced levels of F-actin, DN-myl9 transgenic hearts\ndisplay a coincident reduction in pMyosin levels (suggesting reduced NMII activity) and, by contrast,\ndisplay levels of G-actin that are comparable to the levels in wild-type siblings (Supplementary Fig. 8).\nThese new data suggest that, although actin monomers are readily available in DN-myl9 transgenic\nhearts, they are not being polymerized into filaments to the same degree as in wild-type siblings (p. 11),\npossibly due to the reduced NMII activity in DN-myl9 hearts. Although we did not have a Utr(ABD) line\navailable for these studies, we hope that our G-actin staining approach will help to address the\nreviewer’s queries.\nFigure 4:\nThe authors claim that transplantation addresses the cell-intrinsic versus tissue-scale role of actomyosin;\nhowever, the transplantation leads to large groups of mutant cells rather than the salt-pepper labeling\nthat would allow for a more precise evaluation. Using plasmid injection (rather than transplantation)\nmay be more revealing of whether neighboring wild-type cells influence the shape of the mutant cells or\nnot.\nWe appreciate Reviewer #3’s astute observation regarding the potential influence of neighboring cells\non the shape of donor-derived, transgene-expressing cells. However, in our experiments, we were\npleased to be able to achieve a wide variety of clone sizes (ranging from 1 donor-derived cell to 33\ndonor-derived cells), and not just large groups of donor-derived cells (Reviewer Fig. 3). Furthermore,\nwe found no correlation between clone size and any of the morphometrics tested (For OC cells: apical\nsurface area: R2 = 0.02, p = 0.32; apicobasal length: R2 = 0.01, p = 0.36; apical circularity: R2 = 0.04, p =\n0.12. For IC cells: apical surface area: R2 < 0.01, p = 0.91; apicobasal length: R2 = 0.03, p = 0.39; apical\ncircularity: R2 = 0.04, p = 0.3). Even so, we have performed the analysis of transgene-injected F0\nembryos, as suggested by Reviewer #3 (and Reviewer #1), and we have incorporated these data into our\nrevised manuscript (Supplementary Fig. 10; p. 12). Satisfyingly, we have found that transgene-\nexpressing OC cardiomyocytes in injected F0 embryos exhibited planar expansion defects similar to the\ndefects found in the donor-derived OC cardiomyocytes in our blastomere transplantation experiments\n(Fig. 3 and Supplementary Fig. 10; p. 12). As mentioned above (Reviewer #1, comment #1d), we noted\nsubstantial variation in the expression levels of transgene expression in our injected embryos, which is\ncommon in F0 zebrafish embryos (e.g. Fisher et al., Nature Protocols, 2006). Because we did not\nobserve this variation in the mosaic hearts generated by transplantation, we generally feel that the\ntransplantation approach is a more controlled strategy for mosaic analysis.\nReviewer Figure 3. Clone sizes from Tg(myl7:DN-myl9-eGFP) into wild-type transplantation\nexperiments.\nViolin plots are as in Fig. 3j. Individual datapoints for wild-type host-derived cells have been removed;\nindividual datapoints for DN-myl9 donor-derived cells are colored and sized based on clone size. Note\nthat cells from different clone sizes, ranging from 1 donor-derived cell to 33 donor-derived cells, are\ndistributed broadly along the y-axis.\nFigure 5:\nThe findings regarding the total levels of F-actin remain unclear. A more detailed explanation and\njustification for the quantitative comparison would be helpful.\nWe are unsure of exactly which elements of our original Figure 5 are being addressed by Reviewer #3 in\nthis comment. However, we suspect that Reviewer #3 may share Reviewer #1’s curiosity about the\nreduced F-actin levels observed in myh6 mutant and tbx5a mutant hears. As described above (Reviewer\n#1, minor comment #3), we have incorporated the examination of G-actin levels into our revised\nmanuscript (Supplementary Figs. 12 and 13). Specifically, we have found that myh6 mutants and tbx5a\nmutants exhibit reduced levels of F-actin, together with G-actin levels that are similar to wild-type\n(Supplementary Figs. 12 and 13). As in DN-myl9 transgenics, these data suggest that availability of actin\nmonomers is normal in myh6 mutants and in tbx5a mutants, but that actin modifiers are not properly\nexecuting the normal polymerization of actin filaments in these hearts (pp. 13, 14, 20).\nFigure 8:\nIncluding an illustration or phase diagram that summarizes the effects of various actomyosin and Tbx\nperturbations on apical-basal length and apical surface area would greatly enhance the clarity and the\nworking model.\nWe thank Reviewer #3 for the suggestion to include an additional schematic in our revised manuscript.\nWe have added a new figure (Supplementary Fig. 11; pp. 13, 14, 21, 22, 31, 32) that compares the mean\napical surface areas and mean apicobasal lengths of OC and IC cardiomyocytes at different\ndevelopmental stages and with different genotypes. Altogether, it nicely illustrates the divergence of\nwild-type OC and IC cell morphologies between 37 and 48 hpf, as well as the differences in OC cell\nmorphologies found in myh6 mutants and tbx5a mutants at 48 hpf. While we are unsure whether this\nschematic incorporates all of the information that Reviewer #3 had in mind, we feel that it will be useful\nfor the readers of our revised manuscript."}, {"round": 2, "reviewer_comments": {"reviewer_1": "Whilst the authors have not completed all of the recommended revisions made, they have provided sound justification for why certain experiments were not, or could not, be performed. However, the revision experiments they have completed have enhanced the work and improved the manuscript overall. I also raised concerns over some technical aspects (such as measurements in X-Y versus X-Z) and the response was thorough, clear and alleviated my concerns.\nIn sum, they have addresses sufficient concerns that I now consider the manuscript suitable for publicaiton.", "reviewer_2": "The authors did a reasonable job of responding to critiques and the manuscript is improved by the addition of some data and clarification in the text. I have no further issues and can recommend publication.", "reviewer_3": "The authors have addressed my concerns and the manuscript has significantly improved."}, "author_response": ""}]

Decision letter