
==== Front
Nat Commun
Nat Commun
Nature Communications
2041-1723
Nature Publishing Group UK London

39289341
52340
10.1038/s41467-024-52340-3
Article
Distinct mechanisms regulate ventricular and atrial chamber wall formation
http://orcid.org/0009-0004-9409-8762
Albu Marga 123
Affolter Eileen 123
http://orcid.org/0000-0002-5423-7295
Gentile Alessandra 1236
Xu Yanli 123
http://orcid.org/0000-0001-5672-3791
Kikhi Khrievono 234
http://orcid.org/0009-0008-8410-5550
Howard Sarah 123
http://orcid.org/0000-0001-8013-5906
Kuenne Carsten 5
Priya Rashmi 1237
http://orcid.org/0000-0002-5592-9680
Gunawan Felix 1238
http://orcid.org/0000-0002-0382-0026
Stainier Didier Y. R. didier.stainier@mpi-bn.mpg.de

123
1 https://ror.org/0165r2y73 grid.418032.c 0000 0004 0491 220X Max Planck Institute for Heart and Lung Research, Department of Developmental Genetics, Bad Nauheim, Germany
2 https://ror.org/031t5w623 grid.452396.f 0000 0004 5937 5237 German Centre for Cardiovascular Research (DZHK), Partner Site Rhine-Main, Bad Nauheim, Germany
3 https://ror.org/04ckbty56 grid.511808.5 Cardio-Pulmonary Institute (CPI), Bad Nauheim, Germany
4 grid.4372.2 0000 0001 2105 1091 Flow Cytometry Service Group, Max Planck for Heart and Lung Research, Bad Nauheim, Germany
5 https://ror.org/0165r2y73 grid.418032.c 0000 0004 0491 220X Bioinformatics Core Unit (BCU), Max Planck Institute for Heart and Lung Research, Bad Nauheim, Germany
6 grid.13097.3c 0000 0001 2322 6764 Present Address: MRC Centre for Neurodevelopmental Disorders, King’s College, London, UK
7 https://ror.org/04tnbqb63 grid.451388.3 0000 0004 1795 1830 Present Address: Francis Crick Institute, London, UK
8 https://ror.org/00pd74e08 grid.5949.1 0000 0001 2172 9288 Present Address: Institute of Cell Biology, University of Münster, Münster, Germany
17 9 2024
17 9 2024
2024
15 815919 1 2024
29 8 2024
© The Author(s) 2024
2024
https://creativecommons.org/licenses/by/4.0/ Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/.
Tissues undergo distinct morphogenetic processes to achieve similarly shaped structures. In the heart, cardiomyocytes in both the ventricle and atrium build internal structures for efficient contraction. Ventricular wall formation (trabeculation) is initiated by cardiomyocyte delamination. How cardiomyocytes build the atrial wall is poorly understood. Using longitudinal imaging in zebrafish, we found that at least 25% of the atrial cardiomyocytes elongate along the long axis of the heart. These cell shape changes result in cell intercalation and convergent thickening, leading to the formation of the internal muscle network. We tested factors important for ventricular trabeculation including Nrg/ErbB and Notch signaling and found no evidence for their role in atrial muscle network formation. Instead, our data suggest that atrial cardiomyocyte elongation is regulated by Yap, which has not been implicated in trabeculation. Altogether, these data indicate that distinct cellular and molecular mechanisms build the internal muscle structures in the atrium and ventricle.

Albu et al. show that muscle cells in the cardiac atrium construct structures through heterogeneous cell behaviors that are different, at the cellular and molecular levels, from those involved during ventricular chamber maturation.

Subject terms

Organogenesis
Morphogenesis
Experimental organisms
https://doi.org/10.13039/501100004189 Max-Planck-Gesellschaft (Max Planck Society) issue-copyright-statement© Springer Nature Limited 2024
==== Body
pmcIntroduction

The vertebrate heart consists of distinct chambers: the atria receive the blood from the circulation whereas the ventricles pump blood out of the heart into the circulation. Both the atrial and ventricular muscle walls are composed of complex inner structures important for the function of each chamber. However, it remains unclear whether these internal muscle structures in the different chambers are built through similar or distinct processes.

Cardiac ventricles form complex inner muscular networks known as trabeculae, which increase tissue mass and contraction force, thereby playing critical roles in cardiac function1,2. Cardiac atria form a similarly complex muscular network wherein a large crista terminalis (terminal crest) splits into thinner parallel pectinate muscles and Bachmann’s bundle. Together, these structures play important roles in action potential propagation and chamber contraction1–9. Since cardiac malformations are the most prevalent form of congenital defects10, how the embryonic heart develops to achieve its architecture has been under intense investigation. In addition, abnormal atrial architecture can lead to serious symptoms including arrhythmias3,7,8,11,12.

Atrial and ventricular cardiomyocytes (CMs) are known to have different cellular and molecular profiles as they differentiate13–17. In zebrafish embryos, atrial CMs initially appear more squamous while ventricular CMs appear more cuboidal15,18,19. At the molecular level, atrial and ventricular CM specification and differentiation are regulated in part by different pathways15. For example, the BMP pathway promotes atrial CM specification19,20 whereas the FGF pathway promotes ventricular CM specification21–23. In addition, atrial and ventricular CMs cluster separately in scRNA-sequencing datasets14,24–27, as they express different genes such as MYH6 in atrial CMs and MYH7 in ventricular CMs in fish and mammals15,25,28–30. Nevertheless, how these cellular and molecular differences affect morphogenetic mechanisms remains unexplored.

In the ventricle, the onset of trabeculation has been extensively investigated2,31–53. Trabeculae appear when a subset of the ventricular CMs, which initially form a polarized epithelium, delaminate to form multicellular clusters in the cardiac lumen. This delamination process requires Nrg/ErbB and Notch signaling2,32–34,40–49,52,53. Furthermore, blood flow/cardiac contractility is required for ventricular trabeculation, at least in part by promoting nrg2a expression in the endocardium46,54,55.

However, the cellular and molecular factors involved in atrial wall formation are unclear. Studies in chick have shown that as the atrium develops, inner wall CMs become distinct from outer wall CMs by expressing genes involved in action potential generation and propagation such as Nav1.4 and Cx40, and by becoming more proliferative7,56. Furthermore, a clonal study in zebrafish suggests that up to 7 days post fertilization (dpf), atrial CMs display a squamous and round morphology, whereas at 14 dpf, the atrium appears to consist of a webbed network of rod shaped CMs57. However, the exact mechanisms involved in atrial morphogenesis remain unclear. Here, we use the zebrafish model to investigate this process as it allows for high-resolution longitudinal imaging51.

Results

A subset of atrial cardiomyocytes elongates in the direction of blood flow

In zebrafish, a contracting heart tube is visible by 24 h post fertilization (hpf)58, after which the heart loops to form the atrium and ventricle, which are separated by the atrioventricular canal (AVC)2. The heart then undergoes several morphogenetic events including AV valve formation starting around 56 hpf59, ventricular trabeculation starting around 60 hpf49, and atrial chamber morphogenesis57.

To determine the onset of atrial morphogenesis, we longitudinally imaged atrial development, and documented changes in cell behaviors that might lead to internal muscle structure formation. Taking advantage of the Airyscan imaging modality (Fig. 1a), we obtained clear 3D images of the atrium (Fig. 1b, b’; Supplementary Movie 1). To investigate atrial CM behavior, we imaged zebrafish larvae every 24 h, from 100 to 148 hpf, and observed a subset of CMs gradually elongating over this period (Fig. 1c–e’). To determine when cell elongation begins, we segmented atrial CMs to obtain their 3D shape index – a parameter describing cell shape in 3D - which revealed that a few atrial CMs start to elongate at 76 hpf, and that these shape changes become more prominent with time (Fig. 1f, g); at 100 hpf, approximately 15% of the atrial CMs are elongating, and at 124 hpf approximately 25% (Fig. 1g). During this cell elongation process, atrial CMs orient bidirectionally along the long axis of the heart (Fig. 1b’, h–h”’). These data show that atrial CMs undergo a cell behavior that is distinct from those taking place during ventricular trabeculation, i.e., CM apical constriction and delamination51, or from the previously hypothesized mechanism that drives atrial morphogenesis, i.e., division, budding and branching57.Fig. 1 A subset of atrial cardiomyocytes gradually elongate along the long axis of the heart.

a Schematic of 3D longitudinal imaging. Created with BioRender.com, released under a Creative Commons Attribution-NonCommercial-NoDerivs 4.0 International license. b, b’ 3D image processing used throughout the study. 3D airyscan images of a 100 hpf larva showing both the atrium and ventricle (b) and only the atrium (b’) after 3D cropping in Imaris; CM membranes shown in cyan (myl7:EGFP-Hsa.HRAS) and CM nuclei in yellow (myl7:H2B-mScarlet); magenta square and arrow indicate cropped region; the solid line in b’ is the longest line through one atrial CM, and the dashed lines show how the angle (relative to the AVC) of the highlighted atrial CM (yellow dashed outline) was measured. c–e’ 3D airyscan longitudinal imaging of the atrium at 100, 124, and 146 hpf; CM membranes shown in cyan (myl7:EGFP-Hsa.HRAS) and CM nuclei in yellow (myl7:H2B-mScarlet); one elongating CM shown in 3D (magenta). f 3D quantification of atrial CM shape at 76, 100, and 124 hpf (n = 43 atrial CMs from 6 hearts at 76 hpf, and n = 44 atrial CMs from 6 hearts at 100 and 124 hpf; each data point represents one atrial CM; average of 8 CMs per heart; ordinary one-way ANOVA with Tukey’s multiple comparison test). g Percentage of elongating atrial CMs at 76, 100, and 124 hpf (n = 6 hearts for all time points; each data point represents one heart; average of 20 CMs per heart; Kruskal-Wallis test with Dunn’s multiple comparison test). h–h”, Atrial CM angle measured at 76, 100, and 124 hpf (n = 43 atrial CMs from 6 hearts at 76 hpf, and n = 44 atrial CMs from 6 hearts at 100 and 124 hpf; each data point represents one atrial CM; average of 8 CMs per heart; Kruskal-Wallis test with Dunn’s multiple comparison test; 76 vs 100 hpf p-value = 0.32*10−4, 76 vs 124 hpf p-value = 0.67*10−7). Error bars are mean ± SD.

Atrial cardiomyocyte elongation leads to cell intercalation

We hypothesized that the observed atrial CM elongation drives tissue architecture changes important for internal muscle structure formation. To test this hypothesis, we imaged larval hearts that mosaically express a cytoplasmic marker in atrial CMs, an approach that enables reliable tracking of individual cells (Fig. 2a–d). We then segmented neighboring and elongating CMs in 3D, prior to and during cell elongation. To improve visualization, we created both opaque (Fig. 2a’–d’) and transparent (Fig. 2a”–d”) 3D surfaces. Through this method, we observed that cell elongation leads to cell intercalation, as revealed by the overlap of the mTagBFP- (white surfaces) and mTagBFP+ (magenta surfaces) atrial CMs (Fig. 2a–d”’). To understand how these beating CMs remain attached while intercalating, we examined the localization of the CM-specific cadherin, N-cadherin, in the region of cell intercalation (Fig. 2a”’–d”’). As atrial CMs elongate and intercalate, N-cadherin relocalizes from a continuous distribution on their lateral membranes to a punctate distribution on their apical and basal membranes (Fig. 2a”’–d”’; Supplementary Fig. 1a–b”) similar to what was observed during ventricular CM delamination60. We also noticed that while elongating, CMs form apicobasal N-cadherin-based adhesions, whereas the CMs that maintain their round configuration maintain their lateral N-cadherin based adhesions (Supplementary Fig. 1a–g). In addition, unlike compact layer ventricular CMs, some atrial CMs detached from their neighbors, as revealed by an actin ring (Supplementary Fig. 1h, h’) devoid of N-cadherin (Supplementary Fig. 1i, i’), possibly corresponding to the previously described regions of myocardial absence in the mature atrium57. In summary, we observed that atrial CM elongation leads to regional multilayering via cell intercalation, accompanied by changes in intercellular adhesion.Fig. 2 Atrial cardiomyocyte elongation leads to cell intercalation and convergent thickening.

a–d 3D airyscan imaging of the same larva every 12 h from 82 to 118 hpf; CM membranes shown in white (myl7:mCherry-CAAX) and mosaic CM cytoplasmic expression in magenta (myl7:mTagBFP2). a’–d” 3D segmentation of two elongating and intercalating CMs reconstructed with opaque (a’–d’) and transparent (a”–d”) surfaces, revealing cell intercalation; orange arrow points to the segmented CMs; squares and dashed lines indicate cross-section region. a”’–d”’, Cross-sections through elongating atrial CMs; dashed lines outline the two intercalating CMs; N-cadherin shown in cyan (cdh2:cdh2-EGFP); lat–lateral adhesion; ap–apical adhesion, ba – basal adhesion; (a–d”’) all 3D surfaces of the mTagBFP2+ CM shown in magenta and all 3D surfaces of the mTagBFP2- CM in white. e, f Outer surface views of 3D airyscan images of the same atrium at 76 hpf and 7 dpf; CM membranes shown in cyan (myl7:EGFP-Hsa.HRAS) and CM nuclei in yellow (myl7:H2B-mScarlet). e’, f’ Inner surface views of the same 3D airyscan images at 76 hpf and 7 dpf; orange arrows point to CMs in the inner ridges.

Convergent thickening leads to internal muscle structures in the atrium

We then asked whether this cell intercalation-driven multilayering, also known as convergent thickening61, builds the muscle structures inside the atrial wall. We created 3D surfaces of the atrial myocardium before (Fig. 2e, e’) and after (Fig. 2f, f’) CM elongation and observed ridges on the inner surface of 7 dpf atria. We thus hypothesized that the generation of these ridge-like structures in the atrium constitutes the first step in the formation of internal muscle structures, which are clearly present at 14 dpf (Fig. 3a, a’)57.Fig. 3 Elongated cardiomyocytes form the inner atrial muscle structures.

a–c 3D surface rendering of a 14 dpf fixed atrium; outer layer myocardium shown in white and inner layer myocardium in cyan. a’–c’ 3D confocal image of the atrium from which the surface rendering was created; CM membranes shown in white for outer layer CMs and in cyan for inner layer CMs; all CM nuclei are shown in yellow.

To test the hypothesis that the elongating CMs build these complex atrial muscle structures, we first segmented them from the rest of the myocardium at 14 dpf (Fig. 3a–c; Supplementary Movie 2). We observed that the inner muscle structures of the atrium were entirely composed of elongated CMs (Fig. 3a’–c’). We then used mosaic labeling to track the same atrial CMs from 124 hpf (Supplementary Fig. 2a–c) to 14 dpf (Supplementary Fig. 2a’-c’) and observed that elongating CMs present at 124 hpf form the internal muscle structures at 14 dpf. Furthermore, using this mosaic labeling in combination with longitudinal imaging, we determined and compared the shape index of mTagBFP+ atrial CMs at 124 hpf and 14 dpf. Through this analysis, we observed that a subset of round atrial CMs had become elongated by 14 dpf, leading to a high F-ratio value (Supplementary Fig. 2d). However, elongated atrial CMs never reverted to the round configuration and maintained their shape (Supplementary Fig. 2e). And while all the CMs in the inner layer of 14 dpf atria were elongated, the outer layer contained both elongated and round CMs (Supplementary Fig. 2f). Altogether, these data indicate that CM elongation and intercalation build the internal muscle structures in the zebrafish atrium.

We then investigated whether atrial CM elongation correlates with changes in myofibril organization. We noticed that at 14 dpf, myofibrils of the inner atrial muscle network, which is composed of elongated CMs, are all oriented in the same direction, in contrast to the myofibrils of the outer layer CMs, which display a stochastic arrangement (Supplementary Fig. 3a–c’). We also tracked the myofibrils of the elongating atrial CMs and observed that prior to cell elongation, they display a stochastic orientation (Supplementary Fig. 3d, d’). However, during CM elongation, these myofibrils start aligning in the direction of cell elongation and become significantly thicker (Supplementary Fig. 3d–g). Altogether, these data suggest that atrial CM elongation might regulate myofibril organization, and thus possibly cardiac contraction. Therefore, we hypothesized that atrial CM elongation helps increase contraction forces. We tested this hypothesis by first measuring the atrial ejection fraction before (74 hfp) and after (124 hpf) CM elongation (Supplementary Fig. 3h–i’) and found a significant increase during this time (Supplementary Fig. 3j). Altogether, these data suggest that atrial CM elongation, and associated myofibril maturation, are important to increase atrial contraction forces.

Membrane protrusions drive atrial cardiomyocyte elongation

We next aimed to uncover the cellular processes that underlie atrial CM elongation. High-resolution imaging of the atrial CM membranes prior to cell elongation revealed small protrusions (Fig. 2e), leading us to hypothesize that atrial elongation is an active process driven by membrane protrusion formation. To test this hypothesis, we labeled the membrane protrusions of a few CMs through the mosaic overexpression of PH-Akt1-tdTomato-PEST, a tagged membrane-binding domain of zebrafish Akt1 (Fig. 4a). [AKT1 is known to localize to the leading front of migrating cells62,63, thereby enabling better visualization of membrane protrusions]. We tracked PH-Akt1-tdTomato-PEST+ atrial CMs and observed again their bidirectional elongation and the formation of membrane protrusions that appeared similar to lamellipodia and filopodia (Fig. 4b–d’)64–66. Lamellipodia are large membrane protrusions conventionally thought to be the primary drivers of cell migration, whereas filopodia are smaller protrusions that sense the environment and direct movement64–66. To test whether membrane protrusion formation drives atrial CM elongation, we overexpressed a dominant negative (DN) version of IRSp53 in CMs by using a Tg(UAS:IRSp53DN-RFP) line67 crossed to the Tg(myl7:GAL4) line68. IRSp53DN inhibits actin filament formation in protrusions67,69,70, and upon CM specific IRSp53DN overexpression, the proportion of elongating atrial CMs (Fig. 4e–g) and their orientation (Fig. 4e, f, h–h”) were significantly affected. In addition, we pharmacologically inhibited Rac1, a GTPase required for lamellipodia formation, and observed a significant reduction in atrial CM elongation (Fig. 4i–k), but not orientation (Fig. 4i, j, l–l”). These data are consistent with the role of lamellipodia in driving rather than directing cell migration, contrary to filopodia.Fig. 4 Atrial cardiomyocytes elongate through membrane protrusion formation.

a Schematic of the membrane protrusion labeling of a few CMs, i.e., mosaic expression of the PH domain of Akt1 in CMs. Created with BioRender.com, released under a Creative Commons Attribution-NonCommercial-NoDerivs 4.0 International license. b–d’ 3D airyscan imaging at 76, 100, and 124 hpf; CM membranes shown in white (myl7:EGFP-Hsa.HRAS) and PH-Akt1 localization in magenta (myl7:PH-Akt1-tdTomato-PEST). e, f 3D confocal images of the atrium from control sibling and IRSp53DN overexpressing (myl7:gal4; UAS:IRSp53DN-RFP) larvae at 124 hpf; CM membranes shown in white (myl7:EGFP-Hsa.HRAS). g Percentage of elongating atrial CMs in 124 hpf control sibling and IRSp53DN overexpressing larvae (n = 9 control and n = 8 IRSp53DN; each data point represents one heart; two-tailed Mann-Whitney test). h–h”, Atrial CM angle measured in 124 hpf control siblings and IRSp53DN overexpressing larvae (n = 109 atrial CMs from 9 control hearts and n = 102 atrial CMs from 8 IRSp53DN overexpressing hearts; each data point represents one atrial CM; two-tailed Mann-Whitney test). i, j 3D confocal images of the atrium from DMSO-treated (i) and Rac1 inhibitor-treated (j) larvae at 124 hpf; CM membranes shown in cyan (myl7:EGFP-Hsa.HRAS) and CM nuclei in yellow (myl7:H2B-mScarlet). k Percentage of elongating atrial CMs in DMSO- or Rac1 inhibitor-treated larvae at 124 hpf (n = 7 DMSO and n = 6 Rac1 inhibitor; each data point represents one heart; two-tailed Mann-Whitney test). l–l”, Atrial CM angle measured in 124 hpf DMSO-treated and Rac1 inhibitor-treated larvae (n = 109 atrial CMs from 7 DMSO hearts and n = 105 atrial CMs from 6 Rac1 inhibitor hearts; each data point represents one heart; two-tailed Mann-Whitney test). Error bars are mean ± SD.

Membrane protrusion formation and stabilization are regulated by cell contractility, which depends upon non-muscle myosin activity66. To test the role of non-muscle myosin activity in CM elongation, we overexpressed constitutively active (CA) and DN forms of the non-muscle myosin MYL9 in CMs using the myl7 promoter. Overexpressing MYL9CA significantly increased atrial CM elongation, whereas overexpressing MYL9DN reduced it (Supplementary Fig. 4a–d). As myl7 is active from 15 hpf71, we tested whether atrial CM elongation could be induced to start prematurely or whether there is a mechanism that prevents premature elongation. Surprisingly, even though MYL9CA was present in CMs from an early stage, as assessed by EGFP expression, atrial CM elongation was only observed from 76 hpf onwards, in both control and MYL9CA overexpressing larvae (Supplementary Fig. 4e–k). However, the number of elongating atrial CMs in both 76 and 100 hpf MYL9CA overexpressing larvae was at least doubled compared with control (Supplementary Fig. 4e-k).

Next, we hypothesized that the formation of small membrane protrusions prior to CM elongation was required for the emergence of larger oriented protrusions. To test this hypothesis, we determined the average number of membrane protrusions per CM in 76 hpf atria of wild-type, CM specific MYL9DN overexpressing (Supplementary Fig. 5a–c, Supplementary Table 1), and IRSp53DN overexpressing (Supplementary Fig. 5d–f, Supplementary Table 1) larvae. No reduction in the number of small membrane protrusions was observed in MYL9DN overexpressing larvae (Supplementary Fig. 5a–c, Supplementary Table 1), suggesting that CM specific MYL9DN overexpression does not impair atrial CM elongation by inhibiting the formation of small protrusions. In contrast, IRSp53DN overexpression appeared to reduce the number of small membrane protrusions (Supplementary Fig. 5e, Table 1), perhaps leading to the observed impairment in both CM elongation and orientation (Fig. 4e–h”). Altogether, these data indicate that atrial CM elongation is an active process driven by membrane protrusion formation.

Atrial morphogenesis is not regulated by factors important for ventricular trabeculation

Given the importance of the Notch and Nrg/ErbB signaling pathways during various aspects of cardiac morphogenesis including ventricular wall formation51, we aimed to test whether they were also involved in atrial wall formation. We first searched for Notch signaling pathway activity in the atrium during atrial CM elongation, but observed no Notch reporter expression in the atrium between 100 and 148 hpf (Supplementary Fig. 6a–c). Consistent with these data, the number of elongating atrial CMs and the process of atrial convergent thickening did not change significantly after Notch inhibitor treatment between 72 and 124 hpf (Supplementary Figs. 6d–f). Furthermore, we observed no significant differences in the number of elongating CMs or in the formation of inner myocardial ridges in the atrium of 124 hpf erbb2 mutants compared with homozygous wild-type siblings (Supplementary Fig. 6g–i), despite the presence of nrg2a expression in the larval atrium46. We also tested the role of cardiac contractility since it is also required for various aspects of cardiac morphogenesis including ventricular wall formation41,54, but the number of elongating CMs or the formation of inner myocardial ridges in the atrium were not significantly affected by decreased contractility following BDM treatment from 96 to 124 hpf (Supplementary Fig. 6j–l). Lack of atrial CM contractions in amhc/myh6 mutants72 also did not block atrial CM elongation or intercalation, but slightly reduced the percentage of elongating atrial CMs (Supplementary Fig. 6m–o). However, the abnormal expansion of the atrial chamber in BDM-treated larvae and in myh6 mutants could itself lead to atrial CM elongation, and thus single cell resolution approaches will be required to further address the role of atrial CM contractility in their elongation. Altogether, these data indicate that the key regulators of ventricular trabeculation such as the Notch and Nrg/ErbB signaling pathways are not required for atrial morphogenesis.

Atrial morphogenesis is regulated through Yap activity

To uncover the molecular cues underlying atrial CM elongation, which is clearly visible by 100 hpf (Fig. 1), we isolated 48 and 72 hpf atrial CMs through fluorescence activated cell sorting (FACS), and performed bulk RNA-sequencing (Supplementary Fig. 7). Unbiased pathway enrichment analysis of genes significantly upregulated at 72 hpf compared with 48 hpf uncovered genes promoting cell cycle progression (Supplementary Fig. 8a). We searched for the time point at which most atrial CMs enter S-phase by using a 24-hour EdU pulse assay starting at 48, 72, 96, and 120 hpf (Supplementary Fig. 8b). Interestingly, we found the highest proportion of EdU+ atrial CMs at 96 hpf (Supplementary Fig. 8b). Furthermore, we observed a significant increase in the number of atrial CMs at 124 hpf (Supplementary Fig. 8c). Altogether, these data suggest that at 72 hpf, atrial CMs upregulate the expression of genes that promote cell cycle progression and subsequently proliferate. As such, the peaks of atrial CM elongation (Fig. 1f) and atrial CM proliferation (Supplementary Fig. 8c) coincide at 124 hpf.

The Hippo pathway is a well-known regulator of cell proliferation and tissue morphogenesis73, and several of the proliferation genes significantly upregulated at 72 hpf, including pcna, mcm genes, pola2, cdk1, ccna2, trip13, orc5, esco2, and cdkn1a are known Hippo targets74,75 (Supplementary Fig. 8a). During zebrafish cardiac development, Hippo pathway inhibition through lats1 and lats2 inactivation led to an increase in atrial CM numbers76. Thus, we hypothesized that the increase in atrial CM proliferation observed in wild-type larvae after 74 hpf occurs through changes in Hippo pathway activity. Consistent with this hypothesis, we observed expression of Hippo pathway genes including the transcriptional effector genes yap1 and wwtr1 in atrial CMs at both 48 and 72 hpf (Supplementary Fig. 8d). We then used a Yap1 antibody, previously validated during zebrafish cardiac development77,78, and observed immunostaining in the nucleus of approximately 10% of atrial CMs at 76 hpf, which increased to approximately 25% at 100 hpf, before decreasing to approximately 15% at 124 hpf (Supplementary Figs. 8e–f’). However, nuclear Yap1 immunostaining was not clearly observed in elongating atrial CMs (6 hearts examined at 124 hpf) (Supplementary Fig. 8g, g’).

We then tested whether Yap1 was required for atrial morphogenesis. We incubated larvae from 72 to 124 hpf with K-975 or IWR-1, two inhibitors previously used in zebrafish79–81. K-975 blocks the interaction between Yap and its co-activator Tead82, and IWR-1 is a tankyrase inhibitor that can sequester Yap inside the cytoplasm83. Both K-975 and IWR-1 treatments reduced atrial CM elongation (Fig. 5a–h”). Although IWR-1 can also inhibit the Wnt/β-catenin signaling pathway, we did not observe Wnt/β-catenin reporter activity in atrial CMs (Supplementary Fig. 9a–c), and overexpression of the Wnt/β-catenin pathway inhibitor Axin1 in atrial CMs did not affect atrial CM elongation significantly (Supplementary Fig. 9d–f).Fig. 5 Hippo pathway inhibition affects atrial cardiomyocyte elongation.

a–d 3D airyscan images of atria from 124 hpf larvae treated from 72 to 124 hpf with DMSO (a, c), K-975 (b), or IWR-1(d); CM membranes shown in cyan (myl7:EGFP-Hsa.HRAS) and CM nuclei in yellow (myl7:H2B-mScarlet). e, g 3D quantification of atrial CM shape in 124 hpf larvae treated from 72 to 124 hpf with DMSO, K-975, or IWR-1 (n = 41 atrial CMs from 4 DMSO hearts, n = 36 atrial CMs from K-975 hearts, n = 52 atrial CMs from 5 DMSO hearts, and n = 58 atrial CMs from 5 IWR-1 hearts; each data point represents one atrial CM; average of 10 CMs per heart two-tailed Mann-Whitney test (e); two-tailed unpaired Student’s t-test p-value = 0.48*10-4 (g)). f–f”, h–h”, Atrial CM angle measured in 124 hpf larvae treated from 72 to 124 hpf with DMSO, K-975, or IWR-1 (n = n = 41 atrial CMs from 4 DMSO hearts, n = 36 atrial CMs from 4 K-975 hearts, n = 49 atrial CMs from 5 DMSO hearts, and n = 59 atrial CMs from 5 IWR-1 hearts; each data point represents one atrial CM; average of 10 CMs per heart; two-tailed Mann-Whitney tests). Error bars are mean ± SD.

At the cellular level, atrial CM elongation is an active process driven by cytoskeletal changes (Fig. 4). To investigate whether the Hippo pathway is required upstream or downstream of these cytoskeletal changes, we treated MYL9CA overexpressing larvae and control siblings with K-975 or IWR-1 and observed a rescue in the number of elongating atrial CMs in both treatment conditions (Supplementary Fig. 10a–g). However, K-975 or IWR-1 treatments did not appear to have an effect on the ability of atrial CMs to form small membrane protrusions (Supplementary Fig. 10h–k, Supplementary Table 1). Altogether, these data suggest that atrial CM elongation is regulated, at least in part, by the Hippo pathway, and that the cytoskeletal changes important for CM elongation, but not for small membrane protrusion formation, occur downstream of the Hippo pathway.

Hippo pathway inhibition negatively affected atrial CM elongation, therefore we tested whether this inhibition would also affect cardiac function (Supplementary Fig. 3). K-975 and IWR-1 treatments blocked the increase in myofibril thickness and alignment observed in control larvae (Supplementary Fig. 11a–f). Analysis of the inner atrial myocardial surface of DMSO- versus K-975- or IWR-1-treated larvae revealed the loss of the complex ridges at 124 hpf (Supplementary Fig. 11h–j). Furthermore, the atrial ejection fraction (Supplementary Fig. 11g), but not the heart rate (Supplementary Fig. 11k), was significantly reduced upon both treatments, suggesting that the decreased in atrial CM intercalation and in myofibril organization affects atrial chamber contraction.

Yap1 establishes a cell heterogeneity important for atrial morphogenesis

To investigate how Yap1 regulates atrial CM elongation at the single cell level, we took advantage of a DN form of zebrafish Yap1 fused to a nuclear localization signal (NLS)84,85. We cloned this NLSYap1DN construct downstream of a heat shock-inducible promoter (hsp70l:lox-TagBFP-STOP-lox-NLS-Yap1DN-t2a-mCherry, abbreviated hsp70l:LSL-NLSYap1DN), and injected the resulting plasmid into myh6:creERT2 embryos which allowed for atrial CM-specific expression upon cre-mediated recombination (Fig. 6a–c). Yap1DN overexpression, visualized by mCherry expression, was specifically present in atrial CMs in a mosaic fashion upon tamoxifen treatment and heat shock (Fig. 6c). Notably, quantification of the percentage of elongating CMs that were wild-type (TagBFP+) or Yap1DN positive (mCherry+) revealed more elongating CMs that overexpressed Yap1DN (Fig. 6d), suggesting that CMs with impaired Yap activity have a higher propensity to elongate. Thus, global inhibition of Yap activity reduced the number of elongating atrial CMs, while CMs with impaired Yap activity have a higher propensity to elongate. To investigate this apparent discrepancy, we looked more closely at atrial CM proliferation and observed that global inhibition of Yap activity also reduced atrial CM numbers (Supplementary Fig. 12a, b). To investigate the relationship between CM division and elongation, we incubated larvae from 72 to 124 hpf with the DNA replication inhibitor Aphidicolin, which resulted in a reduction in atrial CM numbers (Supplementary Fig. 12c), consistent with previous data86,87. However, this treatment did not affect the absolute number of elongating atrial CMs, thereby leading to an increase in their percentage (Supplementary Fig. 12d). These data lead us to suggest that after 76 hpf the atrial myocardium becomes heterogeneous with at least two types of atrial CMs, round Yap1+ CMs and elongating Yap1- CMs, and that Yap1 regulates atrial CM elongation independently of its effect on cell division. To test this model, we generated a stable hsp70l:LSL-NLSyap1DN transgenic line using the same construct previously used for the mosaic experiment (Fig. 6a–d). Global overexpression of DNyap1 at early developmental stages using this new line caused cardiac looping defects (Fig. 6e, f), consistent with previous studies of early loss of Yap1 function in zebrafish85,88,89. We then overexpressed DNYap1 in all atrial CMs, after recombination with the myh6:creERT2 transgene, and observed a significant decrease in the number of elongating atrial CMs at 124 hpf (Fig. 6g–i), as previously observed when inhibiting Yap1 function pharmacologically (Fig. 5). Altogether, these data indicate that Yap1 is involved in atrial CM elongation.Fig. 6 Yap modulates atrial cardiomyocyte elongation.

a Schematic of mosaic Yap1DN overexpression in CMs. b, c 3D confocal images of 144 hpf atria from control mosaic labeled (b) (creERT2-, TagBFP+) and Yap1DN mosaic overexpressing (c) (creERT2+, mCherry+) larvae; CM membranes shown in cyan (myl7:EGFP-Hsa.HRAS), control mosaic labeling in magenta (hspl70:LSL-NLSyap1DN, myh6:creERT2-), and Yap1DN mosaic labeling in yellow (hspl70:LSL-NLSyap1DN, myh6:creERT2+). d Percentage of elongating CMs from control labeled or Yap1DN overexpressing CMs (n = 8 hearts, 15 control labeled CMs; n = 8 hearts, 27 YAP1DN overexpressing CMs). e schematic of Yap1DN overexpression using the stable transgenic line. f Percentage of left, centered, and right oriented hearts at 30 hpf from uninjected, water injected, and cre mRNA injected hspl70:LSL-NLSyap1DN embryos (n = 10 uninjected controls, n = 24 water injected controls, and n = 23 cre mRNA injected embryos). g, h 3D airyscan images of 144 hpf atria from hspl70:LSL-NLSyap1DN control larvae (g) and hspl70:LSL-NLSyap1DN; myh6:CreERT2 siblings (h); CM membranes shown in cyan (myl7:EGFP-Hsa.HRAS); larvae were heat shocked once daily from 48 hpf and kept in Tamoxifen from 60 to 144 hpf, refreshed once daily. i, Percentage of elongating atrial CMs at 144 from hspl70:LSL-NLSyap1DN control larvae and hspl70:LSL-NLSyap1DN; myh6:CreERT2 siblings (n = 9 Cre- controls and n = 14 Cre+ siblings, two-tailed Mann-Whitney test). Error bars are mean ± SD.

Distinct mechanisms drive atrial and ventricular muscle structure formation

Since Hippo pathway activity appears to be required for atrial morphogenesis, and Taz/Wwtr1 is important during ventricular wall formation90, we hypothesized that the Hippo pathway was regulating both atrial and ventricular muscle structure formation. To test this hypothesis, we imaged the atrium of wwtr1 mutants and homozygous wild-type siblings at 124 hpf and observed no atrial morphogenesis defects (Fig. 7a–c). Conversely, K-975, IWR-1, or Rac1 inhibitor treatment did not block the onset of trabeculation (Fig. 7d–h). Together these data suggest that the onset of atrial and ventricular muscle structure formation occurs through distinct cellular and molecular mechanisms (Fig. 7i).Fig. 7 Distinct mechanisms regulate atrial and ventricular muscle structure formation.

a, b 3D airyscan images of atria from homozygous wild-type siblings and wwtr1 mutants at 124 hpf; CM membranes shown in cyan (myl7:EGFP-Hsa.HRAS). c Percentage of elongating atrial CMs in 124 hpf homozygous wild-type siblings and wwtr1 mutants (n = 11 wild types, n = 9 mutants; each data point represents one heart; two-tailed Mann-Whitney test). Error bars are mean ± SD. d–g 2D confocal planes of ventricles from 72 hpf larvae treated from 35 to 72 hpf with DMSO, K-975, IWR-1, or Rac1 inhibitor; CM membranes shown in cyan (myl7:EGFP-Hsa.HRAS) and nuclei in yellow (myl7:H2B-mScarlet). h Number of trabecular units in 72 hpf larvae treated from 35 to 72 hpf with DMSO, K-975, IWR-1, or Rac1 inhibitor (n = 5 DMSO, n = 7 K-975, n = 8 IWR-1, n = 6 Rac1 inhibitor; each data point represents one heart; ordinary one-way ANOVA with Dunnett’s multiple comparison test); red arrows point to elongating CMs; white asterisks indicate trabecular units. Error bars are mean ± SD. i Schematic summarizing the cellular and molecular processes involved in ventricular (left) and atrial (right) wall morphogenesis (manually rendered using Adobe Illustrator).

Discussion

Our high-resolution 3D live imaging revealed that starting at early larval stages, a subset of atrial CMs extend membrane protrusions leading to their elongated appearance. This CM elongation leads to cell intercalation and convergent thickening of the atrial myocardium, ultimately resulting in the formation of the inner muscle network.

Atrial CM elongation could be an indirect effect of chamber ballooning36,91 or it could represent an active process. Our observations and manipulations of atrial CM protrusive activity indicate that it is an active process. These atrial CMs elongate along the long axis of the heart and in both directions, i.e., towards the AVC and the sinoatrial node. Thus, these specialized cardiac populations92,93 could provide signals to stimulate atrial CM protrusive activity and/or directionality. Along these lines, we tested the role of Wnt/β-catenin activity, which is present in the AVC and sinoatrial node92,93, in atrial CM elongation but failed to see an effect. Interestingly, only a subset of atrial CMs elongate raising important questions as to how they are selected and what limits their number. Furthermore, not all elongating atrial CMs end up in the inner muscle structures as some remain in the outer layer at least up to 14 dpf. The cellular and molecular mechanisms that select the CMs that form the inner ridges versus those that form the outer layer, as well as the physiological relevance of having an outer layer composed of mixed CM morphology, remain to be investigated. It is possible that maintaining the round atrial CM population is required for the integrity of the tissue as neighboring CMs elongate and intercalate.

While Notch signaling is important in limiting the number of ventricular CMs that delaminate49, it doesn’t appear to play a similar role in atrial CM selection (Supplementary Fig. 6a–f). Conversely, global inhibition of Yap activity reduced atrial CM elongation but it did not stop the onset of ventricular trabeculation. However, inhibiting Yap activity resulted in a reduced number of trabecular units, and wwtr1 mutants also exhibit trabeculation defects90, suggesting that the Hippo signaling pathway is involved in ventricular wall formation, either by modulating CM behavior directly and/or indirectly via the endocardium or epicardium.

Concurrent with atrial CM elongation, we observed atrial CM proliferation; and while these processes appear to be mutually exclusive (as suggested by the data from the Aphidicolin treatments), pharmacological inhibition of Yap activity led to a decrease of both processes. In addition, we observed Yap1 in the nuclei of a subset of atrial CMs, and mosaic Yap1DN overexpression in atrial CMs cell-autonomously promoted their elongation, whereas overexpressing Yap1DN in all atrial CMs using a stable line impaired their elongation. Possible scenarios to explain all these observations include the need for heterogenous Yap1 activation within the atrial myocardium to allow for atrial CM elongation. Yap has been implicated in several different developmental processes including cell proliferation and cell migration94, and more sophisticated tools and approaches will be required to dissect its roles in atrial morphogenesis.

Following CM elongation, cell intercalation and convergent thickening of the atrial myocardium leads to the transformation of a monolayered epithelium into a complex structure. Convergent thickening was originally described to occur within the mesoderm prior to involution during amphibian gastrulation95. While the cellular and molecular basis of convergent thickening remains to be discovered, it was recently reported to be independent of dorsoventral patterning61. Notably, similar to atrial CM intercalation, convergent thickening during gastrulation involves the formation of oriented lamellipodia under the control of cell contractility96, and it will be interesting to investigate how similar convergent thickening is in these two different developmental settings, as well as during other morphological processes.

Ultimately, the ventricular and atrial chambers play distinct roles; while the ventricles need to build strong muscular walls to propel the blood throughout the body, the atria need to provide a structure that can collect blood and propagate the action potential quickly. The elongation of some atrial CMs may provide the means to achieve this quick impulse propagation.

Methods

Zebrafish handling

Zebrafish larvae were raised under standard conditions. All transgenic larvae were presorted for fluorescence. Non-mutant larvae were screened for the presence of developmental defects (e.g., pericardial edema, situs inversus, cardiac size or shape abnormalities) prior to experiments and excluded from the pool if they displayed any.

Adult fish were maintained in 3.5 l tanks at a stock density of 10 fish/l with the following parameters: water temperature: 27–27.5 °C; light:dark cycle: 14:10; pH: 7.0–7.5; conductivity: 750-800 µS/cm. Fish were fed 3–5 times a day, depending on age, with granular and live food (Artemia salina). Health monitoring was performed at least once a year. All procedures performed on animals conform to the guidelines from Directive 2010/63/EU of the European Parliament on the protection of animals used for scientific purposes and were approved by the Animal Protection Committee (Tierschutzkommission) of the Regierungspräsidium Darmstadt (reference: B2/2057). Sample size was calculated to minimize the number of animals used in compliance to the principle of the 3Rs and the European Directive 2010/63/EU.

Zebrafish lines

The transgenic and mutant lines used in this study are: Tg(myl7:EGFP-Hsa.HRAS)s88397, abbreviated myl7:EGFP-Hsa.HRAS; Tg(-0.8myl7:H2B-mScarlet)bns53498, abbreviated myl7:H2B-mScarlet; TgBAC(cdh2:cdh2-EGFP,crybb1:ECFP)zf51799, abbreviated cdh2:cdh2-EGFP; Tg(myl7:mCherry-CAAX)bns748, abbreviated myl7:mCherry-CAAX; Tg(UAS: A53DN-p2a-tagRFP)bns44067, abbreviated IRSp53DN; Tg2(myl7:GAL4)cbg2Tg68, abbreviated myl7:gal4; Tg(myl7:actn3b-EGFP)sd10100, referred to as myl7:actinin3b-EGFP; Tg(myl7:lck-mScarlet)bns561 (this study), abbreviated myl7:lck-mScarlet; Tg(myl7:Hsa.MYL9_T19D,S20D-EGFP)bns33249, abbreviated myl7: MYL9CA-EGFP; Tg(myl7:Hsa.MYL9_T19A,S20A-EGFP)bns33349, abbreviated myl7: MYL9DN-EGFP; Tg(myl7:LIFEACT-GFP)s97440 abbreviated myl7:LIFEACT-GFP; Tg(7XTCF-Xla.Siam:GFP)ia4101, abbreviated 7XTCF-Siam:GFP; Tg(myh7:mCherry-Eco.NfsB)s957102, abbreviated myh7:mCherry-NTR; Tg(-4.5myh6:mTagBFP2)bns629 (this study), abbreviated myh6:mTagBFP2; Tg(hsp70l:LOXP-Luciferase-MYC-STOP-LOXP-Mmu.-Axin1-NLS-dTomato,cryaa:AmCyan)ulm14103, abbreviated hsp70l:LSL-Mmu.Axin1-p2a-NLS-dTomato; Tg(myl7:Cre)sd5543, abbreviated myl7:Cre; Tg(myh6:CreERT2)sd20102, abbreviated myh6:CreERT2; Tg(EPV.Tp1-Mmu.Hbb:Venus-Mmu.Odc1)s940104, abbreviated Tp1-Hbb:Venus-Mmu.Odc1; Tg(hsp70l:loxP-TagBFP-STOP-loxP-NLS-DNyap1-T2A-mCherry)bns729 (this study), abbreviated hsp70l:LSL-NLSyap1DN; erbb2st61 (ref105.); wwtr1bns35 (ref106.); myh6s812 (ref72.).

Plasmid construction

For the membrane localizing myl7:lck-mScarlet construct, the insert was generated through primer annealing (forward primer 5’-GCGGCCATCGAATTGGGATCCCCACCATGGGCTGCGTGTGCAGCAGCAACCCCGAG-3’; reverse primer 5’-GCCCTTGCTCACGGGTACCACCGGTCGCTCGGGGTTGCTGCTGCACACGCAGCCCAT-3’). The Tol2 plasmid -0.8myl7:H2B-mScarlet98 was prepared through BamHI/AgeI restriction digest and the lck insert was inserted. To generate the atrial CM cytoplasmic construct -4.5myh6:mTagBFP2, the mTagBFP2 insert was PCR amplified (forward primer 5’-TCTAGAGCGGCCATCGAATTGGGGATCCCCACCATGGTGTCTAAG-3’; reverse primer 5’-GCCGCCAGTGTGATGGATATCTTAATTAAGCTTGTGCCCCAGTTTGCTAG-3’) from -0.8myl7:mTagBFP2 (gifted from Dr. Rashmi Priya) and inserted into a Tol2 vector containing the -4.5myh6 promoter (gifted from Dr. Thomas Juan). To create the Yap1DN inducible construct hsp70l:loxP-TagBFP-STOP-loxP-NLS-yap1DN-t2A-mCherry, a Tol2 plasmid hsp70l:loxP-TagBFP-STOP-loxP-cxcl18b-t2A-mCherry (gifted from Pinelopi Goumenaki), was digested with XhoI/AgeI to remove the cxcl18b cassette. The pCS2 NLSyap1DN construct85 (gifted from Prof. Virginie Lecaudey) was PCR amplified (forward primer 5’-ATTATACGAAGTTATACCGGTATGGCTCCAAAGAAGAAGCG-3’; reverse primer 5’-GCCCTCTCCACTGCCCTCGAGCCTCAGTCTCTCCTTCTCTA-3’), and inserted in the inducible vector. All fusion experiments were performed using a 5X In-Fusion HD Enzyme Premix (Takara, ST0345).

Zebrafish transgenesis

The -0.8myl7:mTagBFP2 (CM cytoplasmic marker), myl7:PH-Akt1-tdTomato-PEST (CM membrane protrusion marker), and hsp70l:loxP-TagBFP-STOP-loxP-NLSyap1DN-t2A-mCherry (inducible Yap1DN overexpression) Tol2 plasmids were injected together with Tol2 mRNA into one-cell stage wild-type zebrafish embryos for imaging of mosaic hearts (Fig. 4a). The -0.8myl7:lck-mScarlet, -4.5myh6:mTagBFP2, and hsp70l:loxP-TagBFP-STOP-loxP-NLS-yap1DN-t2A-mCherry plasmids were injected together with Tol2 mRNA into one-cell stage wild-type zebrafish embryos which were grown into adulthood. The positive offspring were outcrossed for 2 generations before establishing the stable line.

Immunostaining

Larvae were immunostained as previously described107; however deyolking was skipped to maintain atrial integrity. In brief, larvae were incubated in 0.1% Tricaine/egg water prior to fixation at 4 °C overnight in 4% paraformaldehyde, and then gently washed 3 times with PBS/0.1% Tween; after treatment for 1 hour in PBS/0.1% Tween/5 µg Proteinase K, the larvae were washed 3 times in PBS/1% BSA/1% DMSO/0.5% Triton-X (PBDT), and then blocked for at least 1 hour in PBDT/10% goat serum; primary antibodies were mixed with fresh blocking buffer and incubated with the larvae overnight at 4 °C. The Yap polyclonal antibody was preincubated at 4 °C with zebrafish larvae to improve the signal-noise ratio. Primary antibodies used: anti-Yap1 (rabbit polyclonal, a generous gift from Prof. Virginie Lecaudey, 1:200), anti-eGFP (chicken monoclonal, ab13970 Abcam, 1:200). Secondary antibodies used: Alexa Fluor 488, and Alexa Fluor 647 (1:500, Thermo Fisher Scientific, produced in goat).

EdU staining

Tg(myl7:H2B-mScarlet) larvae were incubated for 24 h in 1-phenyl-2-thiourea (PTU) egg water supplemented with 500 µM EdU and 0.5% DMSO, starting from 48, 72, 96, or 120 hpf. After EdU incubation, the larvae were bathed in 0.2% Tricaine to stop the heart before fixing them in 4% PFA for 2 h at room temperature. Next, the 30 min click-it reaction was performed as per manufacturer’s instructions (Invitrogen, C10340), and confocal imaging was carried out immediately after the washes.

Larval treatments

Zebrafish larvae were incubated in 6-well plates, 6–7 larvae per well, in 4 ml of PTU egg water in which the respective stock reagent was diluted. The incubation time depended on the specific experiment. The solutions were refreshed every 24 h and the larvae were kept in the solution until imaging. The concentrations used in this study were selected based on concentration gradient tests, whereby we used the lowest possible concentration that induced a cardiac phenotype, which was always lower than the concentration that triggered overall toxicity. The control treatments used PTU egg water with the equivalent concentration of DMSO.

The pharmacological reagents used in this study are: NSC23766 (Rac1 inhibitor, incubated from 72 to 124 hpf at 150 µM, or from 35 to 72 hpf at 200 µM, Selleck chemicals S8031), K-975 (Yap/Tead inhibitor, incubated from 72 to 124 hpf at 4 µM, or from 72 to 100 hpf at 4 µM, or from 35 to 72 hpf at 3 µM, Selleck chemicals, E1329), IWR-1 (tankyrase inhibitor, incubated from 72 to 124 hpf at 10 µM, or from 72 to 100 hpf at 10 µM, or from 35 to 72 hpf at 12.5 µM, Selleck chemicals S7086), LY411575 (γ-secretase inhibitor, incubated from 72 to 124 hpf at 2.5 µM, Selleck chemicals S2714), 2,3-butanedione monoxime (BDM) (non-selective myosin ATPase inhibitor, incubated from 96 to 124 hpf at 7.5 mM, Sigma-Aldrich B0753), Aphidicolin (DNA polymerase α inhibitor, incubated from 72 to 124 hpf at 150 µM, Sigma-Aldrich A0781).

To activate the hsp70l promoter, larvae were incubated at 39 °C for 1 h, once daily. To induce CreERT2 nuclear translocation, larvae were treated with 10 µM 4-hydroxytamoxifen (4-OHT) (Sigma H7904, stock diluted in Ethanol) refreshed every 24 h.

Imaging

To avoid pigmentation, for all imaging, embryos/larvae were treated with PTU starting at 24 hpf. To obtain 3D Airyscan images of the entire heart, larvae were mounted in 1% low-melting agarose and 0.2% Tricaine to both anesthetize and stop cardiac contractions during imaging. For longitudinal studies, the larvae were imaged for a maximum of 10 min after which they were carefully recovered in PTU egg water and placed in the incubator to restore cardiac contractions and development. The same larvae were imaged multiple times depending on the experiment (Fig. 1a). For the longitudinal tracking of 124 hpf to 14 dpf hearts, the larvae were imaged live at 124 hpf. The animals were then removed from the agarose and grown in standard conditions up to 14 dpf. At 14 dpf, the same larvae that were imaged at 124 hpf were fixed in 4% PFA and their hearts dissected, mounted in 1% low-melting agarose, and imaged.

3D live images of the entire heart were obtained using the fast Airyscan mode from a Zeiss LSM880 Axio Examiner W Plan-Apochromat 20×/1.0 lens with the Zeiss ZEN software (ZEN 2.3 SP1 FP3 black). Images of fixed or live tissues where only the front of the atrium was required were taken using the confocal mode on a Zeiss LSM880 Axio Examiner W Plan-Apochromat 20×/1.0 lens with the Zeiss Zen software (ZEN 2.3 SP1 FP3 black), or a Zeiss LSM700 Axio Imager W Plan-Apochromat 40×/1.0 dipping lens with the Zeiss ZEN software (ZEN 2011 SP3 black).

Live imaging of the beating heart was obtained by mounting the larvae in 2% agarose without Tricaine to avoid potential arrythmogenic effects. To avoid cardiac contraction fluctuations caused by temperature changes, the embedded larvae were incubated in standard growth conditions for an additional 20 min before imaging using a preheated (27.5 °C) microscope incubator. Cardiac contractions were recorded with a Zeiss Spinning Disk confocal microscope with a 40x/1.1 W lens (ZEN 2.6 blue software), at 5 ms exposure for 20–30 sec. Light intensity and duration were kept at a minimum.

Image processing

All images in the figures are representative images. All airyscan images were processed using ZEN (2.3 SP1 FP3 black), 3D airyscan processing mode (strength auto, 6.0). All 3D renderings, croppings and quantifications were performed using Imaris. The settings used to image the atrium caused a signal saturation in the ventricle (Fig. 1b); therefore, for better visualization, we used the 3D clipping tool in Imaris to exclude the ventricle from all subsequent images, except when the ventricle needed to be analyzed. 3D segmentation of the entire chamber and of single atrial CMs was rendered manually using the myocardial membrane marker as reference and the surface tool to draw, either the entire atrial chamber every 10 planes, or the cell boundaries every plane respectively. Total atrial CM numbers were determined by using the manually rendered atrial chamber surface as a mask to extract the atrial nuclear signal. The spot function was used to segment all atrial nuclei from the extracted signal and to obtain the total number of atrial CMs. The 3D shape index was measured using the 3D surfaces of single atrial CMs and the Ellipsoid-oblate equation available from Imaris (Supplementary Movie 1). To segment the inner atrial muscle structures in 3D, the inner membrane and nuclear signal was manually drawn for every plane to create a 3D mask which was used to extract the outer versus inner layer signal from the original one (Supplementary Movie 2). Both movies were rendered in this way using Imaris.

To obtain the atrial CM angle, the 3D images were first aligned in Imaris. For the alignment, the hearts were positioned such that the AVC CMs, which at these stages have an orientation perpendicular to that of the blood flow in the AV canal, were in a vertical position and the atrium and ventricle were kept parallel to each other (Fig. 1b). 2D images captured from this alignment were obtained using the Imaris Snapshot tool. These 2D images were further analyzed in ImageJ. Only the relatively flat CMs were analyzed and any CM on the curvature of the chamber was left out to avoid artefacts. The longest straight line was drawn through each analyzed CM and the angle of this line in reference to the AVC CM orientation was obtained as a measure of cell angle (Fig. 1b’). To obtain the average number of elongating CMs, the 2D images were first obtained through Imaris pre-alignment and the CMs located away from the curvature were scored for cell shape. To determine the number of small membrane protrusions formed by individual atrial CMs, 3D images were aligned in Imaris, and ImageJ was used to manually count the number of membrane protrusions for each CM. N-cadherin and membrane overlap longitudinal quantification was determined on pre-aligned 2D images. A line was drawn at the same position for each time point and the intensity plot was obtained. The intensity plot revealed regions of high intensity at the membrane junctions and the diameter of these regions was obtained as a measure of the size of N-cadherin or membrane overlap. These diameters were normalized to the first measured time point and then plotted over time.

Myofibril quantifications were performed in ImageJ, on 2D images pre-aligned in Imaris, as described above, and only on myofibrils that were not positioned at the curvature of the chamber. Myofibril angle was measured by drawing a line parallel with the myofibril and the angle of this line in reference to the AVC CM angle was obtained as a measure of myofibril angle. Myofibril thickness was obtained by drawing a line perpendicular to myofibril orientation using the signal from the myl7:actn3b-EGFP Z-band reporter, perpendicular to myofibril orientation. For each wild-type heart, approximately 40 sarcomeres from at least 3 different cells of either round or elongated atrial CMs were averaged to obtain the average sarcomere thickness of the respective cell type. For hearts lacking atrial CM elongation, approximately 60 sarcomeres were measured from at least 6 different cells and averaged to obtain the average sarcomere thickness of that atrium. The percentage of CMs exhibiting nuclear Yap immunostaining was obtained by counting the number of atrial CMs with Yap signal colocalizing with the DAPI signal (nuclear marker) and with the CM membrane marker. This number was then compared to the total DAPI and GFP positive atrial CM nuclei. Heart rate kymographs and ejection fraction were also determined using ImageJ26. Data analysis blinding was only possible for mutant versus wild-type data, as the same person who imaged the other experiments (i.e., the wild-type and the pharmacological ones) also performed them.

Atrial cardiomyocyte isolation and Fluorescence Activated Cell Sorting

To isolate atrial CMs, we used reporter lines that label atrial (myh6:mTagBFP2) and ventricular (myh7:mCherry-NTR) CMs, and a Wnt/β-catenin reporter (7XTCF-Sia:eGFP) that labels the AVC108 and sinus venosus109 CMs (Supplementary Figs. 5a–d’). We were thus able to reliably isolate atrial CMs by sorting the cells positive for mTagBFP2 and negative for eGFP and mCherry (Supplementary Fig 5e-h”).

Approximately 150 hearts at 48 and 72 hpf were extracted through manual dissection and dissociated using the Pierce Cardiomyocyte Isolation Kit (Thermo Fisher Scientific, Catalog# 88281) as previously described98. In brief, hearts were dissected in Dulbecco’s modified Eagle’s medium (DMEM) + GlutaMAX (Thermo Fisher Scientific, Catalog# 10566016) supplemented with 10% FBS and kept on ice throughout the dissociation protocol; the hearts were centrifuged at 4 °C for 5 min at 2300 g; the supernatant was removed and the hearts were washed with 1 ml Hank’s Balanced Salt Solution (HBSS); the tissue was dissociated into single cells by incubating it with 100 µl Enzyme 1 and 5 µl Enzyme 2 from the Pierce Cardiomyocyte Isolation Kit at 30 °C on a shaker set at 300 rpm; 1 ml of DMEM with FBS was added to stop the digestion and removed after centrifugation at 4 °C for 3 min at 800 g; fresh DMEM with FBS was added to resuspend the cells and pass them through a 40 µl-filtered fluorescence-activated cell sorting (FACS) sample tube. 2 µl of DRAQ7™ Dye (Catalog# D15106) was added and incubated in the dark for 10 min at RT. The cell suspension was filtered through a 35 µm nylon Falcon® 5 mL Round Bottom Polystyrene 12 × 75 mm Test Tube (Product# 352235). Cells were sorted using a BD FACSAria™ III (BD Biosciences) or an Invitrogen Bigfoot Spectral Cell Sorter (ThermoFisher Scientific) equipped with a 100 µM nozzle and with 20 psi pressure on the instruments. Live and non-AVC and non–SAN CMs were gated by exclusion of DRAQ7™ Dye using 633 nm excitation paired with 730/45 nm band pass filter and eGFP fluorescence using 488 nm excitation paired with 530/30 nm band pass filter, respectively. To sort atrial CMs (myh6:mTagBFP2+), tagBFP fluorescence was measured with 405 nm excitation paired with 455/14 nm band pass filter or 450/50 nm band pass filter; to sort ventricular CMs (myh7:mCherry-NTR+), mCherry fluorescence was measured with 561 nm excitation paired with 610/20 nm band pass filter. Sorted cells were resuspended in 500 µl Trizol for subsequent RNA extraction. Cytometric data were recorded using FACSDiva software (Version 8.0.1; BD Biosciences), and Sasquatch Software (Version 1.19.2; ThermoFisher Scientific). Data analysis was performed using FlowJo software (Version 10.8, BD Biosciences).

Embryonic zebrafish atrial CMs are low in number and sensitive to dissociation, so in order to get enough cells for RNA-sequencing, a total of 5 sorting sessions were completed to obtain 1000-2000 cells per replicate, 3 replicates per condition. The first two replicates consist of cells sorted using the BD FACSAria™ III and BigFoot and the third replicate consists of cells sorted using the BigFoot only.

For gating (Supplementary Fig. 5e–h”), following exclusion of debris, the cell population was selected (Cells), from which single cells were gated using a FSC-A vs FSC-H parameter (Single cells). Within this single cell population, live cells were selected by exclusion of the DRAQ7™ Dye+ population and eGFP+ AVC and SAN CMs. High intensity mTagBFP2 cells with low or no mCherry signal was used to gate Atrial CMs. Ventricular CMs were gated for mCherry only positive cells as low mCherry signal was present in the atrium, and high mCherry in the ventricle (Supplementary Fig. 5a–d).

Transcriptomic analysis

Total RNA was extracted from the sorted atrial CMs using the miRNeasy micro kit (QIAGEN), combined with on-column DNase digestion (DNase-Free DNase Set, QIAGEN, 217084). Due to low RNA amount quality control steps were skipped. Sequencing was performed on a NextSeq2000 instrument (Illumina) using a P3 flowcell with 1 × 72 bp single end setup. Trimmomatic version 0.39 was employed to trim reads after a quality drop below a mean of Q15 in a window of 5 nucleotides and keeping only filtered reads longer than 15 nucleotides110. Reads were aligned versus Ensembl zebrafish genome version danRer11 (Ensembl release 104) with STAR 2.7.10a111. Alignments were filtered to remove: duplicates with Picard 3.0.0 (Picard: A set of tools (in Java) for working with next generation sequencing data in the BAM format), multi-mapping, ribosomal, or mitochondrial reads. Gene counts were established with featureCounts 2.0.4 by aggregating reads overlapping exons excluding those overlapping multiple genes112. The raw count matrix was normalized with DESeq2 version 1.36.0113. Contrasts were created with DESeq2 based on the raw count matrix. Genes were classified as significantly differentially expressed at average count > 5, multiple testing adjusted p-value < 0.05, and -0.585 <log2FC > 0.585. The Ensemble annotation was enriched with UniProt data (Activities at the Universal Protein Resource (UniProt)).

All downstream analyzes are based on the normalized gene count matrix. For KOBAS114 gene set enrichment analysis, DEGs were split into up/down regulated genes. Significant gene set enrichment was defined by FDR and the top 10 gene sets or enriched pathways were plotted (dashed line: p-value = 0.05). For analysis of only the upregulated genes at 72 hpf, we used KEGG, a pathway enrichment analysis from Metascape115.

Statistics

All statistical analyzes and graphs, excluding the polarity plots and transcriptomic analyzes, were obtained using Graphpad Prism v9.3.1. Before choosing a statistical test, a Gaussian distribution was tested using the D’Agostino–Pearson omnibus and Shapiro-Wilk test for normality. All parametric data, which passed the normality tests, were analyzed using the two-way unpaired Student’s t-test, when two conditions were compared, or ordinary one-way ANOVA paired with Tukey’s or Dunnett’s multiple comparison test when multiple conditions were assessed. All non-parametric data that did not pass the normality test were analyzed using two-way Mann-Whitney’s test when comparing two unpaired conditions, or two-tailed Wilcoxon test when comparing two paired conditions, and Kruskal-Wallis test paired with Dunn’s multiple comparisons test when analyzing multiple conditions. The Student’s t-test was paired with an F test to calculate the F-ratio in Supplementary Fig. 2. All error bars were calculated using mean with SD. The polarity graphs were plotted using R. For all graphs, the p-value cut-off used for significance is 0.05.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Supplementary information

Supplementary Information

Peer Review File

Description of Additional Supplementary Files

Supplementary Movie 1

Supplementary Movie 2

Reporting Summary

Source data

Source Data

Supplementary information

The online version contains supplementary material available at 10.1038/s41467-024-52340-3.

Acknowledgements

We thank Virginie Lecaudey for comments and suggestions as a PhD thesis advisory committee member and for sharing reagents, Thomas Juan and Pinelopi Goumenaki for discussions and sharing reagents, Giulia Boezio for discussions and sharing protocols, David Sedmera and Sara Wickström for discussions and comments on the manuscript, Stefan Guenther for RNA-sequencing and analysis, Simon Perathoner for animal proposal support, and Radhan Ramadass for imaging support. This work was supported by funds from the Max Planck Society to D.Y.R.S.

Author contributions

M.A. designed and performed experiments, analyzed the data, and wrote the manuscript. E.A. performed and analyzed experiments. A.G. and Y.X. helped with heart dissections, protocols, and discussions. K.K. performed FACS sorting. S.H. performed embryonic injections. C.K. helped with transcriptomic analysis. R.P. helped with reagents, transgenic fish lines, and project discussions. F.G. helped with heart dissections and supervised experiments. D.Y.R.S. helped to design experiments and analyze data, supervised the work and wrote the manuscript with input from all the authors.

Peer review

Peer review information

Nature Communications thanks Jinhu Wang and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

Funding

Open Access funding enabled and organized by Projekt DEAL.

Data availability

The bulk RNA-sequencing dataset reported in this paper was deposited in the Gene Expression Omnibus (GEO) database (accession: GSE249149). Source data are provided with this paper.

Competing interests

The authors declare no competing interests.

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
==== Refs
References

1. Sedmera D Pexieder T Vuillemin M Thompson RP Anderson RH Developmental patterning of the myocardium Anat. Rec. 2000 258 319 337 10.1002/(SICI)1097-0185(20000401)258:4<319::AID-AR1>3.0.CO;2-O 10737851
Sedmera, D., Pexieder, T., Vuillemin, M., Thompson, R. P. & Anderson, R. H. Developmental patterning of the myocardium. Anat. Rec. 258, 319–337 (2000).10737851
2. Staudt D Stainier D Uncovering the molecular and cellular mechanisms of heart development using the zebrafish Annu Rev. Genet 2012 46 397 418 10.1146/annurev-genet-110711-155646 22974299
Staudt, D. & Stainier, D. Uncovering the molecular and cellular mechanisms of heart development using the zebrafish. Annu Rev. Genet 46, 397–418 (2012).22974299
3. Wu T-J Role of pectinate muscle bundles in the generation and maintenance of intra-atrial reentry Circulation Res. 1998 83 448 462 10.1161/01.RES.83.4.448 9721702
Wu, T.-J. et al. Role of pectinate muscle bundles in the generation and maintenance of intra-atrial reentry. Circulation Res. 83, 448–462 (1998).9721702
4. Ho SY Anderson RH Sánchez-Quintana D Gross structure of the atriums: more than an anatomic curiosity? Pacing Clin. Electrophysiol. 2002 25 342 350 10.1046/j.1460-9592.2002.00342.x 11990664
Ho, S. Y., Anderson, R. H. & Sánchez-Quintana, D. Gross structure of the atriums: more than an anatomic curiosity? Pacing Clin. Electrophysiol. 25, 342–350 (2002).11990664
5. Dobrzynski H Computer three-dimensional reconstruction of the sinoatrial node Circulation 2005 111 846 854 10.1161/01.CIR.0000152100.04087.DB 15699261
Dobrzynski, H. et al. Computer three-dimensional reconstruction of the sinoatrial node. Circulation 111, 846–854 (2005).15699261
6. Loukas M The clinical anatomy of the crista terminalis, pectinate muscles and the teniae sagittalis Ann. Anat. - Anatomischer Anz. 2008 190 81 87 10.1016/j.aanat.2007.05.002
Loukas, M. et al. The clinical anatomy of the crista terminalis, pectinate muscles and the teniae sagittalis. Ann. Anat. - Anatomischer Anz. 190, 81–87 (2008).
7. Sedmera D Changes in activation sequence of embryonic chick atria correlate with developing myocardial architecture Am. J. Physiol.-Heart Circulatory Physiol. 2006 291 H1646 H1652 10.1152/ajpheart.01007.2005
Sedmera, D. et al. Changes in activation sequence of embryonic chick atria correlate with developing myocardial architecture. Am. J. Physiol.-Heart Circulatory Physiol. 291, H1646–H1652 (2006).
8. Zhao J Three‐dimensional integrated functional, structural, and computational mapping to define the structural “fingerprints” of heart‐specific atrial fibrillation drivers in human heart ex vivo J. Am. Heart Assoc. 2017 6 e005922 10.1161/JAHA.117.005922 28862969
Zhao, J. et al. Three‐dimensional integrated functional, structural, and computational mapping to define the structural “fingerprints” of heart‐specific atrial fibrillation drivers in human heart ex vivo. J. Am. Heart Assoc. 6, e005922 (2017).28862969
9. Guerra A Distinct myocardial lineages break atrial symmetry during cardiogenesis in zebrafish eLife 2018 7 e32833 10.7554/eLife.32833 29762122
Guerra, A. et al. Distinct myocardial lineages break atrial symmetry during cardiogenesis in zebrafish. eLife 7, e32833 (2018).29762122
10. Majumdar U Yasuhara J Garg V In vivo and in vitro genetic models of congenital heart disease Cold Spring Harb. Perspect. Biol. 2021 13 a036764 10.1101/cshperspect.a036764 31818859
Majumdar, U., Yasuhara, J. & Garg, V. In vivo and in vitro genetic models of congenital heart disease. Cold Spring Harb. Perspect. Biol. 13, a036764 (2021).31818859
11. Moe TG Abrich VA Rhee EK Atrial fibrillation in patients with congenital heart disease J. Atr. Fibrillation 2017 10 1612 10.4022/jafib.1612 29250225
Moe, T. G., Abrich, V. A. & Rhee, E. K. Atrial fibrillation in patients with congenital heart disease. J. Atr. Fibrillation 10, 1612 (2017).29250225
12. Collins MM Early sarcomere and metabolic defects in a zebrafish pitx2c cardiac arrhythmia model Proc. Natl Acad. Sci. 2019 116 24115 24121 10.1073/pnas.1913905116 31704768
Collins, M. M. et al. Early sarcomere and metabolic defects in a zebrafish pitx2c cardiac arrhythmia model. Proc. Natl Acad. Sci. 116, 24115–24121 (2019).31704768
13. Bruneau BG Transcriptional regulation of vertebrate cardiac morphogenesis Circulation Res. 2002 90 509 519 10.1161/01.RES.0000013072.51957.B7 11909814
Bruneau, B. G. Transcriptional regulation of vertebrate cardiac morphogenesis. Circulation Res. 90, 509–519 (2002).11909814
14. DeLaughter DM Single-cell resolution of temporal gene expression during heart development Developmental Cell 2016 39 480 490 10.1016/j.devcel.2016.10.001 27840107
DeLaughter, D. M. et al. Single-cell resolution of temporal gene expression during heart development. Developmental Cell 39, 480–490 (2016).27840107
15. Martin KE Waxman JS Atrial and sinoatrial node development in the zebrafish heart J. Cardiovascular Dev. Dis. 2021 8 15 10.3390/jcdd8020015
Martin, K. E. & Waxman, J. S. Atrial and sinoatrial node development in the zebrafish heart. J. Cardiovascular Dev. Dis. 8, 15 (2021).
16. Yao Y Marra AN Yelon D Pathways regulating establishment and maintenance of cardiac chamber identity in zebrafish J. Cardiovascular Dev. Dis. 2021 8 13 10.3390/jcdd8020013
Yao, Y., Marra, A. N. & Yelon, D. Pathways regulating establishment and maintenance of cardiac chamber identity in zebrafish. J. Cardiovascular Dev. Dis. 8, 13 (2021).
17. Gentile A Mechanical forces remodel the cardiac extracellular matrix during zebrafish development Development 2024 151 dev202310 10.1242/dev.202310 38984541
Gentile, A. et al. Mechanical forces remodel the cardiac extracellular matrix during zebrafish development. Development 151, dev202310 (2024).38984541
18. Chi NC Genetic and physiologic dissection of the vertebrate cardiac conduction system PLOS Biol. 2008 6 e109 10.1371/journal.pbio.0060109 18479184
Chi, N. C. et al. Genetic and physiologic dissection of the vertebrate cardiac conduction system. PLOS Biol. 6, e109 (2008).18479184
19. Marques SR Yelon D Differential requirement for BMP signaling in atrial and ventricular lineages establishes cardiac chamber proportionality Dev. Biol. 2009 328 472 482 10.1016/j.ydbio.2009.02.010 19232521
Marques, S. R. & Yelon, D. Differential requirement for BMP signaling in atrial and ventricular lineages establishes cardiac chamber proportionality. Dev. Biol. 328, 472–482 (2009).19232521
20. de Pater E Bmp signaling exerts opposite effects on cardiac differentiation Circ. Res 2012 110 578 587 10.1161/CIRCRESAHA.111.261172 22247485
de Pater, E. et al. Bmp signaling exerts opposite effects on cardiac differentiation. Circ. Res 110, 578–587 (2012).22247485
21. Reifers F Walsh EC Léger S Stainier DYR Brand M Induction and differentiation of the zebrafish heart requires fibroblast growth factor 8 (fgf8/acerebellar) Development 2000 127 225 235 10.1242/dev.127.2.225 10603341
Reifers, F., Walsh, E. C., Léger, S., Stainier, D. Y. R. & Brand, M. Induction and differentiation of the zebrafish heart requires fibroblast growth factor 8 (fgf8/acerebellar). Development 127, 225–235 (2000).10603341
22. Marques SR Lee Y Poss KD Yelon D Reiterative roles for FGF signaling in the establishment of size and proportion of the zebrafish heart Dev. Biol. 2008 321 397 406 10.1016/j.ydbio.2008.06.033 18639539
Marques, S. R., Lee, Y., Poss, K. D. & Yelon, D. Reiterative roles for FGF signaling in the establishment of size and proportion of the zebrafish heart. Dev. Biol. 321, 397–406 (2008).18639539
23. Ilagan R Fgf8 is required for anterior heart field development Development 2006 133 2435 2445 10.1242/dev.02408 16720880
Ilagan, R. et al. Fgf8 is required for anterior heart field development. Development 133, 2435–2445 (2006).16720880
24. Litviňuková M Cells of the adult human heart Nature 2020 588 466 472 10.1038/s41586-020-2797-4 32971526
Litviňuková, M. et al. Cells of the adult human heart. Nature 588, 466–472 (2020).32971526
25. Tucker NR Transcriptional and cellular diversity of the human heart Circulation 2020 142 466 482 10.1161/CIRCULATIONAHA.119.045401 32403949
Tucker, N. R. et al. Transcriptional and cellular diversity of the human heart. Circulation 142, 466–482 (2020).32403949
26. Juan T Control of cardiac contractions using Cre-lox and degron strategies in zebrafish Proc. Natl Acad. Sci. 2024 121 e2309842121 10.1073/pnas.2309842121 38194447
Juan, T. et al. Control of cardiac contractions using Cre-lox and degron strategies in zebrafish. Proc. Natl Acad. Sci. 121, e2309842121 (2024).38194447
27. Farah EN Spatially organized cellular communities form the developing human heart Nature 2024 627 854 864 10.1038/s41586-024-07171-z 38480880
Farah, E. N. et al. Spatially organized cellular communities form the developing human heart. Nature 627, 854–864 (2024).38480880
28. Yutzey KE Gannon M Bader D Diversification of cardiomyogenic cell lineages in vitro Developmental Biol. 1995 170 531 541 10.1006/dbio.1995.1234
Yutzey, K. E., Gannon, M. & Bader, D. Diversification of cardiomyogenic cell lineages in vitro. Developmental Biol. 170, 531–541 (1995).
29. Yelon D Stainier DYR Patterning during organogenesis: genetic analysis of cardiac chamber formation Semin. Cell Developmental Biol. 1999 10 93 98 10.1006/scdb.1998.0278
Yelon, D. & Stainier, D. Y. R. Patterning during organogenesis: genetic analysis of cardiac chamber formation. Semin. Cell Developmental Biol. 10, 93–98 (1999).
30. Meilhac SM Lescroart F Blanpain C Buckingham ME Cardiac cell lineages that form the heart Cold Spring Harb. Perspect. Med 2014 4 a013888 10.1101/cshperspect.a013888 25183852
Meilhac, S. M., Lescroart, F., Blanpain, C. & Buckingham, M. E. Cardiac cell lineages that form the heart. Cold Spring Harb. Perspect. Med 4, a013888 (2014).25183852
31. Mikawa T Borisov A Brown AMC Fischman DA Clonal analysis of cardiac morphogenesis in the chicken embryo using a replication-defective retrovirus: I. formation of the ventricular myocardium Developmental Dyn. 1992 193 11 23 10.1002/aja.1001930104
Mikawa, T., Borisov, A., Brown, A. M. C. & Fischman, D. A. Clonal analysis of cardiac morphogenesis in the chicken embryo using a replication-defective retrovirus: I. formation of the ventricular myocardium. Developmental Dyn. 193, 11–23 (1992).
32. Lee K-F Requirement for neuregulin receptor erbB2 in neural and cardiac development Nature 1995 378 394 398 10.1038/378394a0 7477377
Lee, K.-F. et al. Requirement for neuregulin receptor erbB2 in neural and cardiac development. Nature 378, 394–398 (1995).7477377
33. Meyer D Birchmeier C Multiple essential functions of neuregulin in development Nature 1995 378 386 390 10.1038/378386a0 7477375
Meyer, D. & Birchmeier, C. Multiple essential functions of neuregulin in development. Nature 378, 386–390 (1995).7477375
34. Gassmann M Aberrant neural and cardiac development in mice lacking the ErbB4 neuregulin receptor Nature 1995 378 390 394 10.1038/378390a0 7477376
Gassmann, M. et al. Aberrant neural and cardiac development in mice lacking the ErbB4 neuregulin receptor. Nature 378, 390–394 (1995).7477376
35. Liu J A dual role for ErbB2 signaling in cardiac trabeculation Development 2010 137 3867 3875 10.1242/dev.053736 20978078
Liu, J. et al. A dual role for ErbB2 signaling in cardiac trabeculation. Development 137, 3867–3875 (2010).20978078
36. Moorman AFM Christoffels VM Cardiac chamber formation: development, genes, and evolution Physiological Rev. 2003 83 1223 1267 10.1152/physrev.00006.2003
Moorman, A. F. M. & Christoffels, V. M. Cardiac chamber formation: development, genes, and evolution. Physiological Rev. 83, 1223–1267 (2003).
37. Meilhac SM A retrospective clonal analysis of the myocardium reveals two phases of clonal growth in the developing mouse heart Development 2003 130 3877 3889 10.1242/dev.00580 12835402
Meilhac, S. M. et al. A retrospective clonal analysis of the myocardium reveals two phases of clonal growth in the developing mouse heart. Development 130, 3877–3889 (2003).12835402
38. Gupta V Poss KD Clonally dominant cardiomyocytes direct heart morphogenesis Nature 2012 484 479 484 10.1038/nature11045 22538609
Gupta, V. & Poss, K. D. Clonally dominant cardiomyocytes direct heart morphogenesis. Nature 484, 479–484 (2012).22538609
39. Staudt DW High-resolution imaging of cardiomyocyte behavior reveals two distinct steps in ventricular trabeculation Development 2014 141 585 593 10.1242/dev.098632 24401373
Staudt, D. W. et al. High-resolution imaging of cardiomyocyte behavior reveals two distinct steps in ventricular trabeculation. Development 141, 585–593 (2014).24401373
40. Reischauer S Arnaout R Ramadass R Stainier DYR Actin binding GFP allows 4D in vivo imaging of myofilament dynamics in the zebrafish heart and the identification of Erbb2 signaling as a remodeling factor of myofibril architecture Circulation Res. 2014 115 845 856 10.1161/CIRCRESAHA.115.304356 25228389
Reischauer, S., Arnaout, R., Ramadass, R. & Stainier, D. Y. R. Actin binding GFP allows 4D in vivo imaging of myofilament dynamics in the zebrafish heart and the identification of Erbb2 signaling as a remodeling factor of myofibril architecture. Circulation Res. 115, 845–856 (2014).25228389
41. Samsa LA Cardiac contraction activates endocardial notch signaling to modulate chamber maturation in zebrafish Development 2015 142 4080 4091 10.1242/dev.125724 26628092
Samsa, L. A. et al. Cardiac contraction activates endocardial notch signaling to modulate chamber maturation in zebrafish. Development 142, 4080–4091 (2015).26628092
42. D’Amato G Sequential Notch activation regulates ventricular chamber development Nat. Cell Biol. 2016 18 7 20 10.1038/ncb3280 26641715
D’Amato, G. et al. Sequential Notch activation regulates ventricular chamber development. Nat. Cell Biol. 18, 7–20 (2016).26641715
43. Han P Coordinating cardiomyocyte interactions to direct ventricular chamber morphogenesis Nature 2016 534 700 704 10.1038/nature18310 27357797
Han, P. et al. Coordinating cardiomyocyte interactions to direct ventricular chamber morphogenesis. Nature 534, 700–704 (2016).27357797
44. Jiménez-Amilburu V In Vivo visualization of cardiomyocyte apicobasal polarity reveals epithelial to mesenchymal-like transition during cardiac trabeculation Cell Rep. 2016 17 2687 2699 10.1016/j.celrep.2016.11.023 27926871
Jiménez-Amilburu, V. et al. In Vivo visualization of cardiomyocyte apicobasal polarity reveals epithelial to mesenchymal-like transition during cardiac trabeculation. Cell Rep. 17, 2687–2699 (2016).27926871
45. Li J Single-cell lineage tracing reveals that oriented cell division contributes to trabecular morphogenesis and regional specification Cell Rep. 2016 15 158 170 10.1016/j.celrep.2016.03.012 27052172
Li, J. et al. Single-cell lineage tracing reveals that oriented cell division contributes to trabecular morphogenesis and regional specification. Cell Rep. 15, 158–170 (2016).27052172
46. Rasouli SJ Stainier DYR Regulation of cardiomyocyte behavior in zebrafish trabeculation by Neuregulin 2a signaling Nat. Commun. 2017 8 15281 10.1038/ncomms15281 28485381
Rasouli, S. J. & Stainier, D. Y. R. Regulation of cardiomyocyte behavior in zebrafish trabeculation by Neuregulin 2a signaling. Nat. Commun. 8, 15281 (2017).28485381
47. del Monte-Nieto G Control of cardiac jelly dynamics by NOTCH1 and NRG1 defines the building plan for trabeculation Nature 2018 557 439 445 10.1038/s41586-018-0110-6 29743679
del Monte-Nieto, G. et al. Control of cardiac jelly dynamics by NOTCH1 and NRG1 defines the building plan for trabeculation. Nature 557, 439–445 (2018).29743679
48. Uribe V In vivo analysis of cardiomyocyte proliferation during trabeculation Development 2018 145 dev164194 10.1242/dev.164194 30061167
Uribe, V. et al. In vivo analysis of cardiomyocyte proliferation during trabeculation. Development 145, dev164194 (2018).30061167
49. Priya R Tension heterogeneity directs form and fate to pattern the myocardial wall Nature 2020 588 130 134 10.1038/s41586-020-2946-9 33208950
Priya, R. et al. Tension heterogeneity directs form and fate to pattern the myocardial wall. Nature 588, 130–134 (2020).33208950
50. Yue Y Long-term, in toto live imaging of cardiomyocyte behaviour during mouse ventricle chamber formation at single-cell resolution Nat. Cell Biol. 2020 22 332 340 10.1038/s41556-020-0475-2 32123336
Yue, Y. et al. Long-term, in toto live imaging of cardiomyocyte behaviour during mouse ventricle chamber formation at single-cell resolution. Nat. Cell Biol. 22, 332–340 (2020).32123336
51. Gunawan F Priya R Stainier DYR Sculpting the heart: cellular mechanisms shaping valves and trabeculae Curr. Opin. Cell Biol. 2021 73 26 34 10.1016/j.ceb.2021.04.009 34147705
Gunawan, F., Priya, R. & Stainier, D. Y. R. Sculpting the heart: cellular mechanisms shaping valves and trabeculae. Curr. Opin. Cell Biol. 73, 26–34 (2021).34147705
52. Olejnickova, V. et al. Development of ventricular trabeculae affects electrical conduction in the early endothermic heart. Dev. Dyn.10.1002/dvdy.552 (2022).
53. Grego-Bessa J Nrg1 regulates cardiomyocyte migration and cell cycle in ventricular development Circ. Res 2023 133 927 943 10.1161/CIRCRESAHA.123.323321 37846569
Grego-Bessa, J. et al. Nrg1 regulates cardiomyocyte migration and cell cycle in ventricular development. Circ. Res 133, 927–943 (2023).37846569
54. Fukuda R Mechanical forces regulate cardiomyocyte myofilament maturation via the VCL-SSH1-CFL Axis Developmental Cell 2019 51 62 77 10.1016/j.devcel.2019.08.006 31495694
Fukuda, R. et al. Mechanical forces regulate cardiomyocyte myofilament maturation via the VCL-SSH1-CFL Axis. Developmental Cell 51, 62–77 (2019).31495694
55. Peshkovsky C Totong R Yelon D Dependence of cardiac trabeculation on neuregulin signaling and blood flow in zebrafish Developmental Dyn. 2011 240 446 456 10.1002/dvdy.22526
Peshkovsky, C., Totong, R. & Yelon, D. Dependence of cardiac trabeculation on neuregulin signaling and blood flow in zebrafish. Developmental Dyn. 240, 446–456 (2011).
56. Bressan MC Louie JD Mikawa T Hemodynamic forces regulate developmental patterning of atrial conduction PLOS ONE 2014 9 e115207 10.1371/journal.pone.0115207 25503944
Bressan, M. C., Louie, J. D. & Mikawa, T. Hemodynamic forces regulate developmental patterning of atrial conduction. PLOS ONE 9, e115207 (2014).25503944
57. Foglia MJ Cao J Tornini VA Poss KD Multicolor mapping of the cardiomyocyte proliferation dynamics that construct the atrium Development 2016 143 1688 1696 26989176
Foglia, M. J., Cao, J., Tornini, V. A. & Poss, K. D. Multicolor mapping of the cardiomyocyte proliferation dynamics that construct the atrium. Development 143, 1688–1696 (2016).26989176
58. Stainier DYR Fishman MC Patterning the zebrafish heart tube: acquisition of anteroposterior polarity Developmental Biol. 1992 153 91 101 10.1016/0012-1606(92)90094-W
Stainier, D. Y. R. & Fishman, M. C. Patterning the zebrafish heart tube: acquisition of anteroposterior polarity. Developmental Biol. 153, 91–101 (1992).
59. Gunawan F Gentile A Gauvrit S Stainier DYR Bensimon-Brito A Nfatc1 promotes interstitial cell formation during cardiac valve development in zebrafish Circulation Res. 2020 126 968 984 10.1161/CIRCRESAHA.119.315992 32070236
Gunawan, F., Gentile, A., Gauvrit, S., Stainier, D. Y. R. & Bensimon-Brito, A. Nfatc1 promotes interstitial cell formation during cardiac valve development in zebrafish. Circulation Res. 126, 968–984 (2020).32070236
60. Cherian AV Fukuda R Augustine SM Maischein H-M Stainier DYR N-cadherin relocalization during cardiac trabeculation Proc. Natl Acad. Sci. 2016 113 7569 7574 10.1073/pnas.1606385113 27339140
Cherian, A. V., Fukuda, R., Augustine, S. M., Maischein, H.-M. & Stainier, D. Y. R. N-cadherin relocalization during cardiac trabeculation. Proc. Natl Acad. Sci. 113, 7569–7574 (2016).27339140
61. Shook DR Characterization of convergent thickening, a major convergence force producing morphogenic movement in amphibians eLife 2022 11 e57642 10.7554/eLife.57642 35404236
Shook, D. R. et al. Characterization of convergent thickening, a major convergence force producing morphogenic movement in amphibians. eLife 11, e57642 (2022).35404236
62. Srinivasan S Rac and Cdc42 play distinct roles in regulating PI(3,4,5)P3 and polarity during neutrophil chemotaxis J. Cell Biol. 2003 160 375 385 10.1083/jcb.200208179 12551955
Srinivasan, S. et al. Rac and Cdc42 play distinct roles in regulating PI(3,4,5)P3 and polarity during neutrophil chemotaxis. J. Cell Biol. 160, 375–385 (2003).12551955
63. Fukuda R Proteolysis regulates cardiomyocyte maturation and tissue integration Nat. Commun. 2017 8 14495 10.1038/ncomms14495 28211472
Fukuda, R. et al. Proteolysis regulates cardiomyocyte maturation and tissue integration. Nat. Commun. 8, 14495 (2017).28211472
64. Mattila PK Lappalainen P Filopodia: molecular architecture and cellular functions Nat. Rev. Mol. Cell Biol. 2008 9 446 454 10.1038/nrm2406 18464790
Mattila, P. K. & Lappalainen, P. Filopodia: molecular architecture and cellular functions. Nat. Rev. Mol. Cell Biol. 9, 446–454 (2008).18464790
65. Ridley AJ Life at the Leading Edge Cell 2011 145 1012 1022 10.1016/j.cell.2011.06.010 21703446
Ridley, A. J. Life at the Leading Edge. Cell 145, 1012–1022 (2011).21703446
66. Rottner K Faix J Bogdan S Linder S Kerkhoff E Actin assembly mechanisms at a glance J. Cell Sci. 2017 130 3427 3435 10.1242/jcs.206433 29032357
Rottner, K., Faix, J., Bogdan, S., Linder, S. & Kerkhoff, E. Actin assembly mechanisms at a glance. J. Cell Sci. 130, 3427–3435 (2017).29032357
67. Qi J Apelin signaling dependent endocardial protrusions promote cardiac trabeculation in zebrafish eLife 2022 11 e73231 10.7554/eLife.73231 35225788
Qi, J. et al. Apelin signaling dependent endocardial protrusions promote cardiac trabeculation in zebrafish. eLife 11, e73231 (2022).35225788
68. Mickoleit M High-resolution reconstruction of the beating zebrafish heart Nat. Methods 2014 11 919 922 10.1038/nmeth.3037 25042787
Mickoleit, M. et al. High-resolution reconstruction of the beating zebrafish heart. Nat. Methods 11, 919–922 (2014).25042787
69. Millard TH Structural basis of filopodia formation induced by the IRSp53/MIM homology domain of human IRSp53 EMBO J. 2005 24 240 250 10.1038/sj.emboj.7600535 15635447
Millard, T. H. et al. Structural basis of filopodia formation induced by the IRSp53/MIM homology domain of human IRSp53. EMBO J. 24, 240–250 (2005).15635447
70. Meyen D Dynamic filopodia are required for chemokine-dependent intracellular polarization during guided cell migration in vivo eLife 2015 4 e05279 10.7554/eLife.05279 25875301
Meyen, D. et al. Dynamic filopodia are required for chemokine-dependent intracellular polarization during guided cell migration in vivo. eLife 4, e05279 (2015).25875301
71. Yelon D Horne SA Stainier DYR Restricted expression of cardiac myosin genes reveals regulated aspects of heart tube assembly in zebrafish Developmental Biol. 1999 214 23 37 10.1006/dbio.1999.9406
Yelon, D., Horne, S. A. & Stainier, D. Y. R. Restricted expression of cardiac myosin genes reveals regulated aspects of heart tube assembly in zebrafish. Developmental Biol. 214, 23–37 (1999).
72. Kalogirou S Intracardiac flow dynamics regulate atrioventricular valve morphogenesis Cardiovascular Res. 2014 104 49 60 10.1093/cvr/cvu186
Kalogirou, S. et al. Intracardiac flow dynamics regulate atrioventricular valve morphogenesis. Cardiovascular Res. 104, 49–60 (2014).
73. Ma S Meng Z Chen R Guan K-L The hippo pathway: biology and pathophysiology Annu. Rev. Biochem. 2019 88 577 604 10.1146/annurev-biochem-013118-111829 30566373
Ma, S., Meng, Z., Chen, R. & Guan, K.-L. The hippo pathway: biology and pathophysiology. Annu. Rev. Biochem. 88, 577–604 (2019).30566373
74. Ehmer U Sage J Control of proliferation and cancer growth by the Hippo signaling pathway Mol. Cancer Res 2016 14 127 140 10.1158/1541-7786.MCR-15-0305 26432795
Ehmer, U. & Sage, J. Control of proliferation and cancer growth by the Hippo signaling pathway. Mol. Cancer Res 14, 127–140 (2016).26432795
75. Wang Y MCM6 is a critical transcriptional target of YAP to promote gastric tumorigenesis and serves as a therapeutic target Theranostics 2022 12 6509 6526 10.7150/thno.75431 36185598
Wang, Y. et al. MCM6 is a critical transcriptional target of YAP to promote gastric tumorigenesis and serves as a therapeutic target. Theranostics 12, 6509–6526 (2022).36185598
76. Fukui H Hippo signaling determines the number of venous pole cells that originate from the anterior lateral plate mesoderm in zebrafish eLife 2018 7 e29106 10.7554/eLife.29106 29809141
Fukui, H. et al. Hippo signaling determines the number of venous pole cells that originate from the anterior lateral plate mesoderm in zebrafish. eLife 7, e29106 (2018).29809141
77. Bornhorst D Biomechanical signaling within the developing zebrafish heart attunes endocardial growth to myocardial chamber dimensions Nat. Commun. 2019 10 4113 10.1038/s41467-019-12068-x 31511517
Bornhorst, D. et al. Biomechanical signaling within the developing zebrafish heart attunes endocardial growth to myocardial chamber dimensions. Nat. Commun. 10, 4113 (2019).31511517
78. Peralta M Intraflagellar transport complex B proteins regulate the hippo effector Yap1 during cardiogenesis Cell Rep. 2020 32 107932 10.1016/j.celrep.2020.107932 32698004
Peralta, M. et al. Intraflagellar transport complex B proteins regulate the hippo effector Yap1 during cardiogenesis. Cell Rep. 32, 107932 (2020).32698004
79. Nicenboim J Lymphatic vessels arise from specialized angioblasts within a venous niche Nature 2015 522 56 61 10.1038/nature14425 25992545
Nicenboim, J. et al. Lymphatic vessels arise from specialized angioblasts within a venous niche. Nature 522, 56–61 (2015).25992545
80. Moon, J. & Amatruda, J. F. Biochemical analysis of tankyrase activity in zebrafish in vitro and in vivo. in Wnt Signaling: Methods and Protocols (eds. Barrett, Q. & Lum, L.) 1481, 95–100 (Springer, New York, NY, 2016).
81. Sturtzel C Refined high-content imaging-based phenotypic drug screening in zebrafish xenografts npj Precis. Onc. 2023 7 1 16 10.1038/s41698-023-00386-9
Sturtzel, C. et al. Refined high-content imaging-based phenotypic drug screening in zebrafish xenografts. npj Precis. Onc. 7, 1–16 (2023).
82. Kaneda A The novel potent TEAD inhibitor, K-975, inhibits YAP1/TAZ-TEAD protein-protein interactions and exerts an anti-tumor effect on malignant pleural mesothelioma Am. J. Cancer Res 2020 10 4399 4415 33415007
Kaneda, A. et al. The novel potent TEAD inhibitor, K-975, inhibits YAP1/TAZ-TEAD protein-protein interactions and exerts an anti-tumor effect on malignant pleural mesothelioma. Am. J. Cancer Res 10, 4399–4415 (2020).33415007
83. Kim MK Novel insight into the function of tankyrase (Review) Oncol. Lett. 2018 16 6895 6902 30546421
Kim, M. K. Novel insight into the function of tankyrase (Review). Oncol. Lett. 16, 6895–6902 (2018).30546421
84. Cao X Pfaff SL Gage FH YAP regulates neural progenitor cell number via the TEA domain transcription factor Genes Dev. 2008 22 3320 3334 10.1101/gad.1726608 19015275
Cao, X., Pfaff, S. L. & Gage, F. H. YAP regulates neural progenitor cell number via the TEA domain transcription factor. Genes Dev. 22, 3320–3334 (2008).19015275
85. Miesfeld JB Link BA Establishment of transgenic lines to monitor and manipulate Yap/Taz-Tead activity in zebrafish reveals both evolutionarily conserved and divergent functions of the Hippo pathway Mechanisms Dev. 2014 133 177 188 10.1016/j.mod.2014.02.003 24560909
Miesfeld, J. B. & Link, B. A. Establishment of transgenic lines to monitor and manipulate Yap/Taz-Tead activity in zebrafish reveals both evolutionarily conserved and divergent functions of the Hippo pathway. Mechanisms Dev. 133, 177–188 (2014).24560909
86. Matrone G Laser-targeted ablation of the zebrafish embryonic ventricle: a novel model of cardiac injury and repair Int J. Cardiol. 2013 168 3913 3919 10.1016/j.ijcard.2013.06.063 23871347
Matrone, G. et al. Laser-targeted ablation of the zebrafish embryonic ventricle: a novel model of cardiac injury and repair. Int J. Cardiol. 168, 3913–3919 (2013).23871347
87. Vignes H Extracellular mechanical forces drive endocardial cell volume decrease during zebrafish cardiac valve morphogenesis Developmental Cell 2022 57 598 609 10.1016/j.devcel.2022.02.011 35245444
Vignes, H. et al. Extracellular mechanical forces drive endocardial cell volume decrease during zebrafish cardiac valve morphogenesis. Developmental Cell 57, 598–609 (2022).35245444
88. Mateus R Control of tissue growth by yap relies on cell density and F-actin in zebrafish fin regeneration Development 2015 142 2752 2763 26209644
Mateus, R. et al. Control of tissue growth by yap relies on cell density and F-actin in zebrafish fin regeneration. Development 142, 2752–2763 (2015).26209644
89. Fukui H S1P-Yap1 signaling regulates endoderm formation required for cardiac precursor cell migration in zebrafish Developmental Cell 2014 31 128 136 10.1016/j.devcel.2014.08.014 25313964
Fukui, H. et al. S1P-Yap1 signaling regulates endoderm formation required for cardiac precursor cell migration in zebrafish. Developmental Cell 31, 128–136 (2014).25313964
90. Lai JKH The Hippo pathway effector Wwtr1 regulates cardiac wall maturation in zebrafish Development 2018 145 dev159210 10.1242/dev.159210 29773645
Lai, J. K. H. et al. The Hippo pathway effector Wwtr1 regulates cardiac wall maturation in zebrafish. Development 145, dev159210 (2018).29773645
91. Auman HJ Functional modulation of cardiac form through regionally confined cell shape changes PLOS Biol. 2007 5 e53 10.1371/journal.pbio.0050053 17311471
Auman, H. J. et al. Functional modulation of cardiac form through regionally confined cell shape changes. PLOS Biol. 5, e53 (2007).17311471
92. Minhas R Transcriptome profile of the sinoatrial ring reveals conserved and novel genetic programs of the zebrafish pacemaker BMC Genomics 2021 22 715 10.1186/s12864-021-08016-z 34600492
Minhas, R. et al. Transcriptome profile of the sinoatrial ring reveals conserved and novel genetic programs of the zebrafish pacemaker. BMC Genomics 22, 715 (2021).34600492
93. Abu Nahia K Genomic and physiological analyses of the zebrafish atrioventricular canal reveal molecular building blocks of the secondary pacemaker region Cell. Mol. Life Sci. 2021 78 6669 6687 10.1007/s00018-021-03939-y 34557935
Abu Nahia, K. et al. Genomic and physiological analyses of the zebrafish atrioventricular canal reveal molecular building blocks of the secondary pacemaker region. Cell. Mol. Life Sci. 78, 6669–6687 (2021).34557935
94. Sousa-Ortega A A Yap-dependent mechanoregulatory program sustains cell migration for embryo axis assembly Nat. Commun. 2023 14 2804 10.1038/s41467-023-38482-w 37193708
Sousa-Ortega, A. et al. A Yap-dependent mechanoregulatory program sustains cell migration for embryo axis assembly. Nat. Commun. 14, 2804 (2023).37193708
95. Keller R Danilchik M Regional expression, pattern and timing of convergence and extension during gastrulation of Xenopus laevis Development 1988 103 193 209 10.1242/dev.103.1.193 3197629
Keller, R. & Danilchik, M. Regional expression, pattern and timing of convergence and extension during gastrulation of Xenopus laevis. Development 103, 193–209 (1988).3197629
96. Pfister K Shook DR Chang C Keller R Skoglund P Molecular model for force production and transmission during vertebrate gastrulation Development 2016 143 715 727 10.1242/dev.128090 26884399
Pfister, K., Shook, D. R., Chang, C., Keller, R. & Skoglund, P. Molecular model for force production and transmission during vertebrate gastrulation. Development 143, 715–727 (2016).26884399
97. D’Amico L Scott IC Jungblut B Stainier DYR A Mutation in Zebrafish hmgcr1b Reveals a Role for Isoprenoids in Vertebrate Heart-Tube Formation Curr. Biol. 2007 17 252 259 10.1016/j.cub.2006.12.023 17276918
D’Amico, L., Scott, I. C., Jungblut, B. & Stainier, D. Y. R. A Mutation in Zebrafish hmgcr1b Reveals a Role for Isoprenoids in Vertebrate Heart-Tube Formation. Curr. Biol. 17, 252–259 (2007).17276918
98. Boezio GLM The developing epicardium regulates cardiac chamber morphogenesis by promoting cardiomyocyte growth Dis. Models Mechanisms 2022 16 dmm049571 10.1242/dmm.049571
Boezio, G. L. M. et al. The developing epicardium regulates cardiac chamber morphogenesis by promoting cardiomyocyte growth. Dis. Models Mechanisms 16, dmm049571 (2022).
99. Revenu C Quantitative cell polarity imaging defines leader-to-follower transitions during collective migration and the key role of microtubule-dependent adherens junction formation Development 2014 141 1282 1291 10.1242/dev.101675 24595289
Revenu, C. et al. Quantitative cell polarity imaging defines leader-to-follower transitions during collective migration and the key role of microtubule-dependent adherens junction formation. Development 141, 1282–1291 (2014).24595289
100. Wang J The regenerative capacity of zebrafish reverses cardiac failure caused by genetic cardiomyocyte depletion Development 2011 138 3421 3430 10.1242/dev.068601 21752928
Wang, J. et al. The regenerative capacity of zebrafish reverses cardiac failure caused by genetic cardiomyocyte depletion. Development 138, 3421–3430 (2011).21752928
101. Robertson JK Targeting the Wnt pathway in zebrafish as a screening method to identify novel therapeutic compounds Exp. Biol. Med (Maywood) 2014 239 169 176 10.1177/1535370213514322 24414478
Robertson, J. K. et al. Targeting the Wnt pathway in zebrafish as a screening method to identify novel therapeutic compounds. Exp. Biol. Med (Maywood) 239, 169–176 (2014).24414478
102. Zhang R In vivo cardiac reprogramming contributes to zebrafish heart regeneration Nature 2013 498 497 501 10.1038/nature12322 23783515
Zhang, R. et al. In vivo cardiac reprogramming contributes to zebrafish heart regeneration. Nature 498, 497–501 (2013).23783515
103. Bertozzi A Wu C-C Hans S Brand M Weidinger G Wnt/β-catenin signaling acts cell-autonomously to promote cardiomyocyte regeneration in the zebrafish heart Developmental Biol. 2022 481 226 237 10.1016/j.ydbio.2021.11.001
Bertozzi, A., Wu, C.-C., Hans, S., Brand, M. & Weidinger, G. Wnt/β-catenin signaling acts cell-autonomously to promote cardiomyocyte regeneration in the zebrafish heart. Developmental Biol. 481, 226–237 (2022).
104. Ninov N Borius M Stainier DYR Different levels of Notch signaling regulate quiescence, renewal and differentiation in pancreatic endocrine progenitors Development 2012 139 1557 1567 10.1242/dev.076000 22492351
Ninov, N., Borius, M. & Stainier, D. Y. R. Different levels of Notch signaling regulate quiescence, renewal and differentiation in pancreatic endocrine progenitors. Development 139, 1557–1567 (2012).22492351
105. Lyons DA erbb3 and erbb2 are essential for schwann cell migration and myelination in zebrafish Curr. Biol. 2005 15 513 524 10.1016/j.cub.2005.02.030 15797019
Lyons, D. A. et al. erbb3 and erbb2 are essential for schwann cell migration and myelination in zebrafish. Curr. Biol. 15, 513–524 (2005).15797019
106. Kimelman D Smith NL Lai JKH Stainier DY Regulation of posterior body and epidermal morphogenesis in zebrafish by localized Yap1 and Wwtr1 eLife 2017 6 e31065 10.7554/eLife.31065 29283341
Kimelman, D., Smith, N. L., Lai, J. K. H. & Stainier, D. Y. Regulation of posterior body and epidermal morphogenesis in zebrafish by localized Yap1 and Wwtr1. eLife 6, e31065 (2017).29283341
107. Gunawan F Focal adhesions are essential to drive zebrafish heart valve morphogenesis J. Cell Biol. 2019 218 1039 1054 10.1083/jcb.201807175 30635353
Gunawan, F. et al. Focal adhesions are essential to drive zebrafish heart valve morphogenesis. J. Cell Biol. 218, 1039–1054 (2019).30635353
108. Nguyen CT Langenbacher A Hsieh M Chen J-N The Paf1 complex component Leo1 is essential for cardiac and neural crest development in zebrafish Dev. Biol. 2010 341 167 175 10.1016/j.ydbio.2010.02.020 20178782
Nguyen, C. T., Langenbacher, A., Hsieh, M. & Chen, J.-N. The Paf1 complex component Leo1 is essential for cardiac and neural crest development in zebrafish. Dev. Biol. 341, 167–175 (2010).20178782
109. Burkhard SB Bakkers J Spatially resolved RNA-sequencing of the embryonic heart identifies a role for Wnt/β-catenin signaling in autonomic control of heart rate eLife 2018 7 e31515 10.7554/eLife.31515 29400650
Burkhard, S. B. & Bakkers, J. Spatially resolved RNA-sequencing of the embryonic heart identifies a role for Wnt/β-catenin signaling in autonomic control of heart rate. eLife 7, e31515 (2018).29400650
110. Bolger AM Lohse M Usadel B Trimmomatic: a flexible trimmer for Illumina sequence data Bioinformatics 2014 30 2114 2120 10.1093/bioinformatics/btu170 24695404
Bolger, A. M., Lohse, M. & Usadel, B. Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinformatics 30, 2114–2120 (2014).24695404
111. Dobin A STAR: ultrafast universal RNA-seq aligner Bioinformatics 2013 29 15 21 10.1093/bioinformatics/bts635 23104886
Dobin, A. et al. STAR: ultrafast universal RNA-seq aligner. Bioinformatics 29, 15–21 (2013).23104886
112. Liao Y Smyth GK Shi W featureCounts: an efficient general purpose program for assigning sequence reads to genomic features Bioinformatics 2014 30 923 930 10.1093/bioinformatics/btt656 24227677
Liao, Y., Smyth, G. K. & Shi, W. featureCounts: an efficient general purpose program for assigning sequence reads to genomic features. Bioinformatics 30, 923–930 (2014).24227677
113. Love MI Huber W Anders S Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2 Genome Biol. 2014 15 550 10.1186/s13059-014-0550-8 25516281
Love, M. I., Huber, W. & Anders, S. Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biol. 15, 550 (2014).25516281
114. Xie C KOBAS 2.0: a web server for annotation and identification of enriched pathways and diseases Nucleic Acids Res 2011 39 W316 W322 10.1093/nar/gkr483 21715386
Xie, C. et al. KOBAS 2.0: a web server for annotation and identification of enriched pathways and diseases. Nucleic Acids Res 39, W316–W322 (2011).21715386
115. Zhou Y Metascape provides a biologist-oriented resource for the analysis of systems-level datasets Nat. Commun. 2019 10 1523 10.1038/s41467-019-09234-6 30944313
Zhou, Y. et al. Metascape provides a biologist-oriented resource for the analysis of systems-level datasets. Nat. Commun. 10, 1523 (2019).30944313
