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Hippo signaling determines the number of venous pole cells that originate from the anterior lateral plate mesoderm in zebrafish

  1. Hajime Fukui
  2. Takahiro Miyazaki
  3. Renee Wei-Yan Chow
  4. Hiroyuki Ishikawa
  5. Hiroyuki Nakajima
  6. Julien Vermot
  7. Naoki Mochizuki  Is a corresponding author
  1. National Cerebral and Cardiovascular Center Research Institute, Japan
  2. University of Strasbourg Institute for Advanced Study (USIAS), France
  3. Institut de Génétique et de Biologie Moléculaire et Cellulaire, France
  4. Centre National de la Recherche Scientifique, France
  5. Institut National de la Santé et de la Recherche Médicale, France
  6. Université de Strasbourg, France
  7. Japan Agency for Medical Research and Development (AMED), Japan
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Cite as: eLife 2018;7:e29106 doi: 10.7554/eLife.29106

Abstract

The differentiation of the lateral plate mesoderm cells into heart field cells constitutes a critical step in the development of cardiac tissue and the genesis of functional cardiomyocytes. Hippo signaling controls cardiomyocyte proliferation, but the role of Hippo signaling during early cardiogenesis remains unclear. Here, we show that Hippo signaling regulates atrial cell number by specifying the developmental potential of cells within the anterior lateral plate mesoderm (ALPM), which are incorporated into the venous pole of the heart tube and ultimately into the atrium of the heart. We demonstrate that Hippo signaling acts through large tumor suppressor kinase 1/2 to modulate BMP signaling and the expression of hand2, a key transcription factor that is involved in the differentiation of atrial cardiomyocytes. Collectively, these results demonstrate that Hippo signaling defines venous pole cardiomyocyte number by modulating both the number and the identity of the ALPM cells that will populate the atrium of the heart.

https://doi.org/10.7554/eLife.29106.001

Introduction

The human heart typically has about 2 billion cardiomyocytes (CMs) (Adler and Costabel, 1975; Laflamme and Murry, 2011), which together form the muscle layer of the heart responsible for contraction. The determination of the final number of CMs in the different parts of the heart involves highly coordinated processes of cell fate specification and proliferation during development. Understanding the relative contributions of these processes during the different stages of cardiac morphogenesis, as well as the mechanisms behind them, is one of the long-standing goals of the cardiac development field.

Mammalian heart morphogenesis is best studied in the mouse. Early in mouse development, a bilateral group of cells in the splanchnic mesoderm specifies into cardiac precursor cells (CPCs) (Saga et al., 1999) and forms the first heart field (FHF). Cells of the FHF then extend toward the midline to form a crescent-shaped epithelium, known as the cardiac crescent. Through a series of morphogenetic steps, the cardiac crescent gives rise to a structure called the heart tube (Kelly et al., 2014; Vincent and Buckingham, 2010). CPCs from the secondary heart field (SHF), which are derived from pharyngeal mesoderm, are added to the arterial and venous poles of the heart tube (Cai et al., 2003; Kelly et al., 2001; Waldo et al., 2001). The FHF is believed to give rise mostly to the left ventricle and parts of the atria, whereas the SHF is believed to give rise mostly to the right ventricle, the outflow tract (OFT) and most of the atria (Cai et al., 2003; Galli et al., 2008; Waldo et al., 2001; Zaffran et al., 2004).

The zebrafish has a simpler heart than that of mouse and humans, containing only two chambers. Nevertheless, the successive phases of CM differentiation during development, as well as their associated genetic pathways, are well conserved between zebrafish and other vertebrates (Staudt and Stainier, 2012). Because of the optical clarity and external development of zebrafish embryos, as well as the amenability of zebrafish to genetic manipulation, the zebrafish is an excellent model for the study of cardiac development. In the early stages of zebrafish development, a bilateral group of cells in the anterior lateral plate mesoderm (ALPM) specifies into CPCs, and the region where they reside is termed the heart field (HF) (Fishman and Chien, 1997). In zebrafish, both the FHF and the SHF derive from the ALPM (Mosimann et al., 2015). As in the mouse, the zebrafish FHF forms the initial heart tube while the SHF elongates the heart tube by adding to its arterial and venous poles. LIM domain transcription factor Islet1 (Isl1) marks a subset of SHF cells (de Pater et al., 2009; Witzel et al., 2012) that eventually give rise to the inflow tract (IFT) of the atrium of the mature heart. A second set of SHF cells is positive for Islet family member Islet2b (Isl2b) and for latent TGFβ binding protein 3 (Ltbp3). These cells become the CMs that populate the arterial pole of the heart tube and eventually contribute to the OFT of the ventricle in the mature heart (Zhou et al., 2011; Witzel et al., 2017).

Given that CMs derive from the FHF and SHF cells of the mesoderm, the final CM number in the mature heart depends in part on the specification of mesodermal cells to form CPCs. Nkx2.5, Mef2c, and Hand2 are known to promote CM differentiation in zebrafish (Guner-Ataman et al., 2013; Hinits et al., 2012; Lazic and Scott, 2011; Schindler et al., 2014). Several signaling pathways have been found to act upstream of some of these transcription factors and to restrict the HF at the rostral and caudal boundaries of the ALPM. At the rostral border of the zebrafish ALPM, Tal1 and Etv2, two transcription factors required for vascular and hematopoietic lineage specification, respectively, repress cardiac specification, thereby reducing the number of CMs in the mature heart (Schoenebeck et al., 2007). At the border between the zebrafish ALPM and posterior LPM (PLPM), retinoic acid (RA) signaling from the adjacent forelimb field determines the HF size by restricting the potential differentiation of ALPM cells into heart precursor cells, and thus limits the number of atrial, but not ventricular, cells in the mature heart (Waxman et al., 2008).

In addition to molecular pathways regulating the fate decisions of CPCs, the final number of CMs is also determined by signalling pathways that regulate cell proliferation at each stage of cardiac morphogenesis (de Pater et al., 2009; Jopling et al., 2010; Rochais et al., 2009; Vincent and Buckingham, 2010). One well-characterized signaling pathway is the Hippo signaling pathway, which helps to define the number of cells in a variety of tissues and organs (Zhou et al., 2015). In mammalian cells, the active elements of the Hippo pathway include Ste20-like serine/threonine kinase 1 and 2 (Mst1/2, mammalian orthologs of the fruit fly, Hippo), which phosphorylate Large tumour suppressor kinase 1 and 2 (Lats1/2). Phosphorylated Lats1/2 induce the nuclear export of the transcription factor Yes-associated protein 1 (Yap1) and its paralog WW-domain-containing transcription regulator 1 (Wwtr1), also known as Taz. Lats1/2 inhibits the formation of a complex involving Yap1/Wwtr1 and the TEA domain (TEAD) transcription factors by promoting the nuclear export of Yap1/Wwtr1, thereby repressing the expression of downstream target genes (Zhao et al., 2008).

In CMs of the mouse heart, Hippo signaling has been implicated in cardiac regeneration after myocardial injury (Lin et al., 2014; von Gise et al., 2012; Xin et al., 2013). While Yap1 and Wwtr1 double-null mutations in mice are embryonically lethal before the blastula stage (Nishioka et al., 2009), it has been shown that Nuclear Yap1 induces CM proliferation in the adult and fetal mouse. Furthermore, mice that are depleted of Lats2, Salvador (Salv), or Mst1/2 using CM-specific Cre drivers exhibit a hypertrophic growth due to an increase in CM proliferation (Zhou et al., 2015). Together, these results suggest that Hippo signaling plays a key role in cardiac proliferation in the mouse. However, it is unclear whether Yap1/Wwtr1 are involved in CPC proliferation within the FHF and SHF before the formation of the heart tube. In addition, although Hippo signaling also regulates the expression of genes that are essential for cell specification and differentiation (Zhao et al., 2008; Nishioka et al., 2009), we still do not know whether Hippo signalling plays a role in cardiac cell fate specification.

In the work described here, we sought to examine the role of Hippo signaling in controlling heart cell number beyond its known roles in CM proliferation. Using zebrafish as a model, we examined the role of Hippo signaling at various stages of embryonic development: at the stage when embryos are specifying the HF, at the stage when the heart tube is formed, and in older embryos when heart morphogenesis is largely completed. We demonstrate that Lats1/2-Yap1/Wwtr1-regulated Hippo signaling determines the number of SHF cells in the venous pole that originate from the caudal part of the ALPM. At the molecular level, we show that Yap1/Wwtr1 promote bone morphogenetic protein-2b (bmp2b) expression and induce the phosphorylation of Smad1/5/9 in both hand2- and Isl1-positive cells. Consistently, the absence of lats1/2 leads to increased hand2 expression at the boundary between the ALPM and the PLPM and to an increased number of SHF cells in the venous pole. Together, these findings demonstrate that Hippo signaling restricts the number of CPCs located in the venous pole by suppressing Yap1/Wwtr1-dependent Bmp2b expression and hand2 expression.

Results

Lats1/2 are involved in atrial CMs development

To examine whether Yap1/Wwtr1-dependent transcription determines the CM number during early cardiogenesis, we developed lats1 and lats2 knockout (KO) fish using transcription activator-like effector nuclease (TALEN) techniques. Fish with lats1ncv107 and lats2ncv108 alleles lack 10 bp at Exon 2 and 16 bp at Exon 3, respectively, resulting in premature stop codons due to frameshift mutations (Figure 1—figure supplement 1A). lats1ncv107 KO fish and lats2ncv108 KO fish were viable with no apparent defect (data not shown). However, almost all the lats1ncv107lats2ncv108 double KO (lats1/2 DKO) larvae died before 15 days post-fertilization (dpf) (Figure 1—figure supplement 1B).

We assessed the effect of Lats1/2 depletion on heart development by counting CM number in the atrium and the ventricle of lats1/2 mutant larvae which also contained Tg(myosin heavy chain 6 [myh6]:Nuclear localization signal [Nls]-tagged tdEosFP);Tg(myosin light polypeptide 7 [myl7]:Nls-mCherry). These larvae expressed Nls-tdEosFP in the atrial CMs and Nls-mCherry in the whole CMs (Figure 1A). We found that the number of atrial CMs, but not ventricular CMs, was significantly increased in the lats1wt(wild type)/ ncv107lats2ncv108 embryos and in the lats1ncv107lats2ncv108 embryos at 74 hr post-fertilization (hpf) (Figure 1B,C and Figure 1—source data 1).

Figure 1 with 2 supplements see all
Knockout of lats1/2 genes leads to an increase in the number of atrial, but not ventricular CMs during early development.

(A) Confocal 3D-stack images (at 74 hr post fertilization [hpf]) of the Tg(myh6:Nls-tdEosFP);Tg(myl7:Nls-mCherry) embryos with lats1wt/wtlats2wt/wt (top) and lats1ncv107lats2ncv108 alleles (bottom). Atrial (A) and ventricular (V) cardiomyocytes (CMs) are EosFP-positive cells and EosFP-negative mCherry-positive cells, respectively. Ventral view, anterior to the top. (B, C) Quantitative analyses of the number of atrial (B) and ventricular (C) CMs of the embryos at 74 hpf with alleles indicated at the bottom. Plus (+) and minus (–) signs indicate the wt allele and the allele of ncv107 or ncv108 in lats1 or lats2 genes, respectively. The confocal 3D-stack images are a set of representative images of eight independent experiments. In the graphs, the total number of larvae examined in the experiment is indicated on the top of columns unless otherwise described. *p < 0.05.

https://doi.org/10.7554/eLife.29106.002

To confirm that Hippo signaling is involved in determining the number of CMs, we assessed whether Yap1/Wwtr1-dependent transcription is activated in developing CMs by analyzing two Tead-reporter transgenic lines: first, we used a general Tead reporter line, the Tg(eef1a1l1:galdb-hTEAD2ΔN-2A-mCherry), which expresses human TEAD2 lacking the amino-terminus (1–113 aa) fused with a GAL4 DNA-binding domain under the control of an eukaryotic translation elongation factor 1 alpha 1, like 1 (eef1a1l1) promoter (Fukui et al., 2014); second, we generated a CM-specific Tead reporter line, the Tg(myl7:galdb-hTEAD2ΔN-2A-mCherry), which expresses the same construct under the control of the myl7 promoter. By crossing these fish with Tg(uas:GFP) lines, cells with nuclear-translocated Yap1 or Wwtr1 can be identified through their expression of GFP (Fukui et al., 2014).

We first showed that the lats1/2 double knock-out (DKO) affected Hippo signaling using the general Tead reporter. We found that the Yap1/Wwtr1 reporter was active in the lats1/2 DKO embryos and in the lats1/2 morphants (Figure 1—figure supplement 1C). We next analyzed the activity of the CM reporter at 74 hpf and found that Yap1/Wwtr1-dependent transcription was present in IFT atrial CMs using the CM Tead reporter line (Figure 1—figure supplement 2A).

To identify the stage at which the Tead reporter starts to be active in CMs more precisely, we analyzed the activity of the reporter at an earlier stage. Previous studies have shown that IFT atrial CMs originate from the venous pole of the heart tube (de Pater et al., 2009). By examining the progeny of the Tead reporter line crossed with the Tg(myl7:Nls-mCherry) line at 24 hpf, we found that the Tead reporter was active in CMs located at the venous pole of the heart tube (Figure 1—figure supplement 2B). Importantly, the eef1a1l1 and 2A peptide-driven mCherry expression used for screening purposes is very weak compared to mCherry expression driven by the myl7 promoter and does not affect the intensity analysis. Together, these data suggest that Lats1/2 restrict Yap1/wwtr1-Tead activation during early CM determination.

Lats1/2 determine the number of IFT CMs derived from Isl1-positive SHF cells

Following on from the observation that the Tead reporter is active at 24 hpf in the progeny of SHF cells, we next assessed whether Hippo signaling affects CM number at this stage. Isl1 is a SHF marker because it plays an essential role in the development of CPCs in the SHF, and because its expression delimitates the SHF as early as 24 hpf in zebrafish (Caputo et al., 2015). We generated transgenic fish expressing GFP under the control of an isl1 BAC promoter; the TgBAC(isl1:GFP). The isl1 transgene recapitulated the endogenous isl1 expression patterns (Figure 2—figure supplement 1A,B) and expression was found in the IFT of the atrium at four dpf (Figure 2A,B and Figure 2—source data 1). isl1-promoter-active cells were observed in the endocardium and epicardium at 96 hpf (Figure 2—figure supplement 1C, arrows and arrowheads), but they were absent from the arterial pole, OFT, and ventricular myocardium until four dpf (Figure 2A,B and Figure 2—source data 1). To validate our isl1 reporter further, we made use of Ajuba, a LIM-domain family protein that is known to restrict the number of isl1-positive cells in the SHF (Witzel et al., 2012). Consistently, isl1-promoter-active SHF cells were increased in the ajuba morphants (Figure 2E, Figure 2—figure supplement 1D and Figure 2—source data 1). Together, these results suggest that the TgBAC(isl1:GFP) line recapitulates isl1 expression in vivo.

Figure 2 with 2 supplements see all
The Hippo signaling pathway is involved in the formation of Isl1-positive SHF cells in the venous pole.

(A) Single-scan confocal images (at 96 hpf) of the TgBAC(isl1:GFP);Tg(myl7:Nls-mCherry) embryos. Cells in which both the isl1 and myl7 promoters are active are present in the inflow tract (IFT) cells (arrows) but not in the outflow tract (OFT) cells (square bracket). A, atrium; V, ventricle. Ventral view, anterior to the top. (B) Quantitative analyses of the number of cells with both isl1- and myl7-promoter activities in the IFT and the OFT at 96 hpf (n = 10). (C) Confocal images (at 26 hpf) of the TgBAC(isl1:GFP);Tg(myl7:Nls-mCherry) embryos with the lats1wt/wtlats2wt/wt (upper panels) or lats1wt/ncv107lats2ncv108 alleles (bottom panels). The boxed regions are enlarged in the panels in the fourth (right) column. Yellow arrows indicate both isl1- and myl7-promoter activities in cells in the venous pole. White arrowheads indicate cells with isl1-promoter activity that are in contact with cells in which there is myl7-promoter activity. Confocal 3D-stack images (left two panels) and single-scan images (right two panels). Dorsal view, anterior to the top. (D, E) Quantitative analyses of the number of the isl1-promoter-active SHF cells in the venous pole of the lats1wt/wtlats2wt/wt embryos, the lats1wt/ncv107lats2ncv108 embryos, and the lats1/2 DKO embryos (D), and in embryos shown in Figure 2—figure supplement 1D (E). Both cells with isl1- and myl7-promoter activity and with isl1-promoter activity that were in contact cells with myl7-promoter activity were counted as SHF cells. The confocal 3D-stack images and single-scan (2 μm) images are a set of representative images of at least four independent experiments. **p < 0.01.

https://doi.org/10.7554/eLife.29106.006

We found that a significant number of Tead reporter-positive CMs were positive for Isl1 in the venous pole (Figure 2—figure supplement 2A). Furthermore, the number of both Isl1- and Tead-reporter-positive CMs in the venous pole was significantly increased in the lats1/2 morphants (Figure 2—figure supplement 2B). Making use of the TgBAC(isl1:GFP) line, we next quantified the population of isl1-promoter-positive SHF cells in the lats1/2 DKO embryos at 26 hpf. As expected, the number of isl1-promoter-active cells in the venous pole was significantly increased in both lats1wt/ncv107lats2ncv108 and lats1/2 DKO embryos (Figure 2C,D and Figure 2—source data 1). Consistent with this, the isl1-promoter-active SHF cells were significantly increased in the venous pole of the lats1/2 morphants (Figure 2E, Figure 2—figure supplement 1D and Figure 2—source data 1). To further confirm the importance of Hippo signaling in the SHF, we analyzed embryos expressing a mCherry-tagged dominant-negative form of Yap1/Wwtr1-Tead-dependent transcription (ytip-mCherry) (Fukui et al., 2014) and found that the number of isl1-positive cells were significantly decreased in the venous pole (Figure 2E, Figure 2—figure supplement 1D and Figure 2—source data 1). Together, these results demonstrate that Lats1/2-mediated Hippo signaling is involved in reducing the number of SHF-derived CPCs that contribute to the venous pole.

Lats1/2 determine the number of CMs derived from the hand2-promoter-active CMs

To identify the mechanism of action of Hippo signaling, we sought to identify the gene targets of Yap1/Wwtr1 in early CPC differentiation. We examined whether Yap1/Wwtr1 regulate the expression of transcription factors nkx2.5, hand2, and gata4, all of which are essential for early CPC differentiation (Schoenebeck et al., 2007). qPCR revealed that hand2 mRNA expression was significantly upregulated (Figure 3A and Figure 3—source data 1) and that nkx2.5 and gata4 mRNAs expression was unaffected in the lats1/2 morphants (Figure 3A and Figure 3—source data 1). Whole-mount in situ hybridization (WISH) analyses revealed that the hand2 expression domain, corresponding to the region that gives rise to the heart, was expanded in the lats1wt/ ncv107lats2ncv108 embryos, the lats1/2 DKO embryos, and the lats1/2 morphants at 22 hpf (Figure 3B and Figure 3—figure supplement 1A). These data suggest that Lats1/2 determine the number of atrial CMs by inhibiting Yap1/Wwtr1-dependent hand2 expression.

Figure 3 with 2 supplements see all
Knockout of lats1/2 results in an increase in the number of cells in the venous pole in which both myl7 and hand2 promoters are activated.

(A) Quantitative-PCR analyses of the expression of nkx2.5, hand2, and gata4 mRNAs in the whole embryos at 24 hpf showing the effects of MO injection (n = 4). Relative expression of mRNA in the MO-injected morphants to that of the control is shown. (B) WISH analyses at 22 hpf of lats1wt/wtlats2wt/wt (n = 7), lats1wt/ncv107lats2wt/ncv108 (n = 22), lats1wt/ncv107lats2ncv108 (n = 14) and lats1ncv107lats2ncv108 (n = 6) embryos using an antisense probe for hand2. (C) Confocal 3D-stack images (at 26 hpf) of TgBAC(hand2:GFP);Tg(myl7:Nls-mCherry)-labeled embryos carrying the lats1wt/wtlats2wt/wt (upper panels) or lats1ncv107lats2ncv108 allele (bottom panels). GFP images (left), merged GFP and mCherry images (center), and enlarged images of the boxed regions in the center panels (right) are shown. (D) Quantitative analysis of the number of cells in which both hand2 and myl7 promoters are activated at 26 hpf. (E) Confocal 3D-stack images (at 26 hpf) of the TgBAC(hand2:GFP);Tg(myl7:Nls-mCherry)-labeled embryos containing the lats1wt/wtlats2wt/wt (upper panels, n = 9) or the lats1ncv107lats2ncv108 allele (bottom panels, n = 5) immunostained with the anti-Isl1 antibody (anti-Isl1 Ab). Square brackets denote the SHF cells that are Isl1-positive, both hand2- and myl7-promoter-active cells that are Isl1-positive, and hand2-promoter-active cells that are in contact with myl7-promoter-active cells. pp indicates the pharyngeal pouch, which expresses the hand2-promoter-activated GFP signal. The first, second, third and fourth columns show Isl1 immunostaining, merged images of GFP and Isl1 immunostaining, merged images of Isl1 immunostaining and mCherry labeling, and merged images of all the three (GFP, mCherry, and Isl1 immunostaining), respectively. All of the images are of the dorsal view, anterior to the top. The confocal 3D-stack images and the WISH images are a set of representative images from at least four independent experiments. **p < 0.01.

https://doi.org/10.7554/eLife.29106.010

Overexpression of Hand2 increases the number of SHF-derived CMs in zebrafish (Schindler et al., 2014), so we hypothesized that the increased number of CM in lats1/2 mutants is due to an increase in the number of CPCs in the SHF. We first investigated endogenous hand2 expression during CM development by analyzing TgBAC(hand2:GFP) (Yin et al., 2010), which labels cells in which the hand2 promoter is activated with GFP, and Tg(myl7:Nls-mCherry), which labels CMs and CM progenitors with nuclear-localized mCherry. At 26 hpf, we found that hand2-promoter-active CMs are localized on the anterior side of the growing cardiac tube, corresponding to the venous pole (Figure 3C). In the lats1/2 DKO embryos, the number of hand2-promoter-active CMs was significantly increased in the venous pole (Figure 3C,D and Figure 3—source data 1). Similarly, the number of hand2-promoter-active CMs was increased in the venous pole in the lats1/2 morphants (Figure 3—figure supplement 1B). Furthermore, we found that the population of Isl1-positive SHF cells overlaps with the hand2-promoter-active CMs in the very left-rostral end of the cardiac tube (Figure 3E, brackets). As expected, the population of Isl1-positive SHF cells was increased in the lats1/2 DKO embryos and in the lats1/2 morphants (Figure 3E and Figure 3—figure supplement 1C, brackets). Thus, hand2 is expressed in differentiated CMs and the hand2 expression domain contains SHF cells at 26 hpf.

To confirm the involvement of the Hippo pathway in modulating the hand2 expression domain, we next sought to examine the number of hand2-promoter-active CMs in the DKO mutants of yap1 (Figure 3—figure supplement 2A) and wwtr1 (Nakajima et al., 2017). As expected, we found that the extension of the embryo along the anterior-posterior axis is severely impaired in yap1 and wwtr1 DKO embryos (Figure 3—figure supplement 2B) (Kimelman et al., 2017; Nakajima et al., 2017). When analyzing the hand2-promoter-active cells, we found that their number is greatly reduced (Figure 3—figure supplement 2C). Interestingly, the DKO mutant embryos exhibit cardia bifida, which is also observed in the hand2 mutant (Yelon et al., 2000) (Figure 3—figure supplement 2C,D). Together, these results strongly suggest that the increased CPC number in the lats1/2 mutants is due to an expansion of the hand2 expression domain in the SHF.

hand2-promoter-active cells at the caudal end of the ALPM migrate toward the venous pole of the cardiac tube

In zebrafish, the origin of venous pole CMs is unknown. In amniotes, the venous pole progenitors are located in the most caudal domain of the ALPM (Abu-Issa and Kirby, 2008; Galli et al., 2008). Considering that hand2 is expressed in the zebrafish LPM (Schoenebeck et al., 2007), we hypothesized that hand2-promoter-active cells originate from the caudal end of the ALPM. We performed time-lapse imaging from 14 hpf to 26 hpf to investigate whether hand2-promoter-active cells of the LPM contribute to the venous pole cells. Cell-tracking analysis revealed that the caudal cells of the ALPM migrate toward the venous pole (Figure 4A,B, and Video 1). ALPM cells move toward the posterior of the cardiac disc by 20 hpf, and subsequently move anterior-laterally toward the venous pole of the cardiac tube by 26 hpf (Figure 4A–C). These results indicate that hand2-promoter-active cells in the venous pole differentiate from the caudal ALPM. To confirm that the caudal ALPM cells are incorporated into the IFT atrial CMs, we sought to perform cell-tracking of hand2-promoter-active cells in the caudal region of ALPM cells following photoconversion. To do so, we injected embryos with a plasmid that expresses tdEosFP under the control of hand2 BAC promoter and photoconverted the cells in the caudal region of both sides of ALPM in these embryos. We found that the photoconverted cells were incorporated into the IFT of the atrium and the OFT of the ventricle (Figure 4D). This indicates that the cells of the caudal region of both sides of ALPM can become IFT CMs of the atrium.

Figure 4 with 1 supplement see all
Tead-reporter-active cells in the caudal region of the ALPM move to the venous pole.

(A) Time-sequential 3D-rendered confocal images of the TgBAC(hand2:GFP) embryo from 15.5 hpf to 25.5 hpf (n = 6). Spots of magenta and cyan denote the cells in the caudal part of the left and right ALPM, respectively. Notochord, nc; cardiac disc, cd; cardiac tube, ct. ALPM, cd, and ct are indicated by the blue, yellow, and red broken lines, respectively. (B) Tracking of caudal end hand2-promoter-active ALPM cells from 14 hpf to 26 hpf. The 3D-rendered confocal image of the TgBAC(hand2:GFP) embryo at 26 hpf with the track of cells showing color changes from blue to red according to the time after imaging (14 hpf to 26 hpf). (C) Schematic illustration of the trajectory patterns of the caudal end ALPM cells from 14 hpf to 26 hpf. Magenta and cyan denote the caudal region of the left and right ALPM, respectively. The cells in the caudal region of the left ALPM (magenta) and the right ALPM (cyan) moved to the venous pole (26 hpf) through the SHF region posterior to the cd (20 hpf). (D) Confocal 3D-stack images of the Tg(myl7:EGFP) embryo injected with pTol2-hand2 BAC:tdEosFP at the 12 somite stage (ss) (upper panels) and at 52 hpf (lower panels). Cells from the caudal region of either left ALPM (left four panels, n = 19) or right ALPM (right four panels, n = 18) were photoconverted at 12 ss. The hearts of the photoconverted embryos were imaged at 52 hpf. White squares and white lines indicate the photoconverted area and the outline of the heart, respectively. (E) Confocal 3D-stack images (at 12 ss) of the TgBAC(hand2:GFP);Tg(eef1a1l1:galdb-hTEAD2ΔN-2A-mCherry);Tg(uas:mRFP1) embryo (n = 6). Tead-reporter-active cells were present in the entire ALPM. Images of the ALPM are in dorsal view, anterior to the top. Images of the heart are in ventral view, anterior to the top. The confocal 3D-stack images are a set of representative images from at least five independent experiments.

https://doi.org/10.7554/eLife.29106.014
Video 1
hand2-promoter-active cells in the caudal region of the ALPM move to the venous pole.

Time-lapse recording of 3D-rendered confocal images of the TgBAC(hand2:GFP) embryo from 14 hpf (10 ss) to 26 hpf. Note the migration of the caudal region of both the left ALPM (magenta) and the right ALPM (cyan) toward the venous pole of the cardiac tube. Changes in the colors reflect the tracking time (blue, 0 hr; red, 12 hr). Dorsal view, anterior to the top. The time-lapse movie is a set of representative data from six independent experiments. Video 1 is related to Figure 4A,B.

https://doi.org/10.7554/eLife.29106.016

We next analyzed whether the Tead reporter is active in the hand2 expression domain of the ALPM. We crossed TgBAC(hand2:GFP) lines with general Tead mRFP1 reporter fish. At the 12 somite stage (ss), the Tead-reporter-active cells were found in the entire region of the ALPM and overlapped with hand2-promoter-active cells in ALPM (Figure 4E). We further noticed that the Tead reporter was inactive in the rostral region of the PLPM at 12 ss (Figure 4—figure supplement 1A). These data suggest that Hippo signaling acts upstream of hand2 expression in the ALPM and may play a role in determining CM fate in the ALPM.

Hippo signaling regulates the number of SHF cells from the caudal end of the ALPM

Tead reporter activation in the cells of ALPM prompted us to ask whether Lats1/2-Yap1/Wwtr1 signaling is involved in the proliferation and/or specification of those cells. We examined the proliferation of isl1-promoter-active cells using the EdU incorporation assay and found that the number of isl1-promoter-active and EdU-positive CMs in the lats1/2 morphants was comparable to that of controls (Figure 5A,B). In addition, there was no difference in the number of EdU-positive blood cells and endocardial cells among the two groups (data not shown). Importantly, the timing of EdU incorporation did not affect the results of the proliferation analyses (Figure 5B), suggesting that the increase in the number of isl1-promoter-active SHF cells that resulted from the depletion of Lats1/2 is not caused by cell proliferation after the differentiation of SHF cells from the ALPM.

Figure 5 with 1 supplement see all
Knockout of lats1/2 leads to an increase in the expression of hand2 in the boundary between ALPM and PLPM.

(A) Single-scan confocal images (at 96 hpf) of the TgBAC(isl1:GFP);Tg(myl7:Nls-mCherry) embryos injected with MO and pulsed with EdU from 14 hpf to 26 hpf. Arrows indicate the EdU-incorporated isl1- and myl7-promoter-active cells in the IFT of the atrium. A, atrium; V, ventricle. Ventral view, anterior to the top. (B) The number of EdU-positive isl1-promoter-active CMs among the embryos treated with MO. Embryos pulsed with EdU from 14 hpf to 26 hpf (left two columns) and from 20 hpf to 36 hpf (right two columns). (C) Schematic illustration of gene expression patterns in the LPM of wildtype (WT) embryos at 10 somite stage (ss). Expression domain of tal1, gata4, nkx2.5 and hand2 are depicted as magenta, yellow, green, and blue, respectively. Dorsal view, anterior to the top. (D, F, G) WISH analyses of the embryos at 10 ss using the antisense probes indicated to the left of the panels. (D, F) Genotypes are WT (left panels, n = 8 to 18), lats1/2 DKO (center panels, n = 6 to 13), and yap1/wwtr1 DKO (right panels, n = 5 to 7). (D) Square brackets indicate the gap between hand2-positive regions of ALPM and PLPM. (E) Quantitative measurement of the distance indicated by the brackets in (D) in either the lats1wt/wtlats2wt/wt embryos or the lats1 wt/ncv107lats2 wt/ncv108 embryos and in either the lats1wt/ncv107lats2ncv108 or the lats1/2 DKO embryos. (F) Brackets indicate the tal1-positive rostral end of PLPM in the WT. (G) Genotypes are WT (left panels, n = 4 to 5) and lats1/2 DKO (right panels, n = 3 to 4). Dorsal view, anterior to the top. The single-scan (2 μm) confocal images and in situ hybridization (ISH) images are a set of representative images of at least four independent experiments. **p < 0.01.

https://doi.org/10.7554/eLife.29106.017

We next tested whether Lats1/2 affect SHF cell specification in the ALPM. At 10 ss, the ALPM and the PLPM can be characterized by the expression of gata4, nkx2.5, tal1, and hand2. gata4 labels the multipotent myocardial-endothelial-myeloid progenitors of the ALPM; nkx2.5 is a marker for the ventricular HF; tal1 marks the hematopoietic cell progenitors; hand2 marks both the ALPM and the PLPM at 10 ss (Figure 5C) with a clear gap in between. Interestingly, the gap length of the lats1wt/ncv107lats2ncv108 embryos, the lats1/2 DKO embryos, and the lats1/2 morphants was significantly shorter than that of wildtype embryos (Figure 5D,E, Figure 5—figure supplement 1A and Figure 5—source data 1). We also found that tal1 expression was decreased in the rostral end of the PLPM in the lats1/2 DKO embryos and the lats1/2 morphants (Figure 5F and Figure 5—figure supplement 1B). A similar analysis in yap1/wwtr1 DKO embryos revealed that hand2 expression was decreased in both ALPM and PLPM but that the expression of tal1 was unaffected (Figure 5D,F). Importantly, the expression of gata4 and nkx2.5 was unaffected in lats1/2 DKO embryos and in lats1/2 morphants, suggesting that Hippo signaling mainly affects the hand2 expression domain (Figure 5G, and Figure 5—figure supplement 1C). These observations were consistent with the results of qRT-PCR (Figure 3A).

To confirm the specificity of Hippo action, we examined the expression of etv2, which is a marker for blood-vessel progenitors (Schoenebeck et al., 2007), and of hoxb5b, which is a regulatory molecule of RA signaling in the forelimb field (Waxman et al., 2008) in the LPM. The expression levels of both etv2 and hoxb5b were comparable between the control and the lats1/2 morphants (Figure 5—figure supplement 1C). Collectively, these results suggest that Lats1/2 negatively regulate Yap1/Wwtr1-dependent differentiation of the LPM into the SHF at the boundary between ALPM and PLPM.

Hippo signaling regulates Bmp-dependent smad activation that determines the number of SHF cells in the venous pole

Signaling mediated by Bone morphogenetic proteins (Bmps) affects various contexts of heart development via Smad phosphorylation-dependent transcriptional activation. Bmp-Smad signaling is known to be essential for SHF formation, FHF-derived CM development, endocardium development and epicardium development (Prall et al., 2007; Schlueter et al., 2006; Tirosh-Finkel et al., 2010; Yang et al., 2006). Yap1 is known to promote Bmp2b expression in zebrafish neocortical astrocyte differentiation (Huang et al., 2016) and Bmp2 in mouse endothelial cells (Neto et al., 2018). In the zebrafish embryo, bmp2b, but not bmp4, is expressed in the LPM (Chung et al., 2008). We hypothesized that Yap1/Wwtr1 are involved in bmp2b-dependent signaling during early cardiogenesis. To investigate whether Bmp-Smad signaling is activated in the ALPM, we examined Bmp-dependent transcription using the Tg(BRE:GFP) fish embryos, in which the Bmp-responsive element (BRE) drives GFP expression (Collery and Link, 2011). At 14 hpf, BRE-positive cells were found in the ALPM (Figure 6—figure supplement 1A). Because Bmps induce the phosphorylation of Smad1/5/9 (Smad9 is also known as Smad8) (Heldin et al., 1997), we examined the phosphorylation of Smad1/5/9 in the embryos at 14 hpf using immunohistochemistry. The phosphorylated Smad1/5/9-positive cells were found in the ALPM and eyes (Figure 6A). Phosphorylation of Smad1/5/9 was enhanced in the lats1/2 DKO embryos and decreased in the yap1/wwtr1 DKO embryos (Figure 6A). At 10 ss, bmp2b expression was increased in the ALPM and eyes of the lats1/2 DKO embryos and decreased in the yap1/wwtr1 DKO embryos (Figure 6B). Consistently, bmp2b mRNA was increased in the lats1/2 morphants at 10 ss (Figure 6—figure supplement 1B). Although we could not detect bmp4 in the ALPM in the early ss (data not shown), bmp4 mRNA was increased in the venous pole of the lats1/2 morphants at 26 hpf (Figure 6—figure supplement 1C). These results suggest that Hippo signaling functions upstream of Bmp-dependent Smad activation in the ALPM during early cardiogenesis.

Figure 6 with 3 supplements see all
The Hippo signaling pathway functions upstream of the Bmp-dependent signal that is necessary for Isl1-positive SHF formation.

(A) Confocal 3D-stack images (at 14 hpf) of embryos immunostained with the anti-pSmad1/5/9 Ab. Blue broken lines indicate the phosphorylated Smad1/5/9-positive cells in the ALPM and eyes. Note that the pSmad1/5/9-positive signals in the ALPM and the eyes are increased and decreased in the embryos of lats1/2 DKO and yap1/wwtr1 DKO, respectively (n = 3). Scale bar indicates 60 μm. (B) WISH analyses at the 10 somite stage (ss) of WT (n = 9), lats1/2 DKO (n = 10), and yap1/wwtr1 DKO (n = 8) embryos, using antisense probe for bmp2b. Square brackets indicate the bmp2b-positive ALPM. (C) Confocal 3D-stack images (at 24 hpf) of the Tg(BRE:GFP);Tg(myl7:Nls-mCherry) embryos carrying lats1wt/wtlats2wt/wt (upper panels) and lats1wt/ncv107lats2ncv108 alleles (bottom panels). Square brackets highlight the GFP-positive myl7-promoter-active cells in the venous pole. Note that the numbers of GFP-positive myl7-promoter-active cells are increased in the venous pole. (D) Quantitative analyses of the numbers of the both BRE-active GFP-positive and myl7-promoter-active mCherry-positive cells in the lats1wt/wtlats2wt/wt embryos and in either the lats1wt/ncv107lats2ncv108 embryos or the lats1/2 DKO embryos. (E) Confocal 3D-stack images (at 26 hpf) of the TgBAC(isl1:GFP);Tg(myl7:Nls-mCherry) control embryos (uninjected, upper panels) and embryos injected with 100 pg of smad7 mRNA (bottom panels). Square brackets indicate the region of isl1-promoter-active GFP-positive SHF cells in the venous pole. (F) Quantitative analyses of the number of isl1-promoter-active SHF cells in the venous pole. Note that overexpression of smad7 mRNA leads to a decrease in the number of isl1-promoter-active SHF cells in the venous pole. (G) Confocal 3D-stack images (at 26 hpf) of the TgBAC(isl1:GFP);Tg(myl7:Nls-mCherry) embryos treated with DMSO (upper panels) or DMH1 (10 μM, lower panels) constructed between 14 hpf and 26 hpf. Square brackets indicate the isl1-promoter-active SHF cells in the venous pole. (H) Quantitative analyses of the number of isl1-promoter-active SHF cells and of the number of both GFP-negative and myl7-promoter-active mCherry-positive cells. Note that DMH1 treatment decreases the number of isl1-promoter-active SHF cells in the venous pole but does not affect the number of GFP-negative and myl7-promoter-active CMs. All images in this figure are in dorsal view, anterior to the top. The confocal 3D-stack images and ISH images are a set of representative images of at least three independent experiments. **p < 0.01.

https://doi.org/10.7554/eLife.29106.020

By analyzing the BRE reporter, we found that the number of Bmp signal-active cells marked by GFP in the venous pole was increased in the lats1wt/ncv107lats2ncv108 embryos and/or the lats1/2 DKO embryos, as well as in the lats1/2 morphants at 24 hpf (Figure 6C,D, Figure 6—figure supplement 2A and Figure 6—source data 1). Immunohistochemistry revealed that the number of phosphorylated Smad1/5/9-positive and hand2-promoter-active cells was also increased at the venous pole of the lats1/2 morphants at 26 hpf (Figure 6—figure supplement 2B). These results suggest that Yap1/Wwtr1 promote bmp2b expression and subsequent Bmp signaling and that Lats1/2 restrict Yap1/Wwtr1-dependent Bmp signaling leading to the formation of the proper venous pole.

We further investigated whether Bmp-Smad activation promotes hand2 expression. We made use of Smad7, an inhibitory-Smad that blocks Bmp-Smad signaling by interacting with activated Bmp type I receptors and preventing the downstream activation of receptor-regulated Smads (Souchelnytskyi et al., 1998). We first showed that overexpression of smad7 mRNA caused dorsalization, demonstrating that Smad7 is essential for proper Bmp-Smad signaling, using 200 pg of mRNA (Figure 6—figure supplement 3A,B). Interestingly, the injection of lower concentration of mRNA (100 pg) led to a dorsalization phenotype in only 10% of the injected embryos. Therefore, the remaining embryos without dorsalization phenotype were used to assess heart development (Figure 6—figure supplement 3A,B). The non-dorsalized embryos looked healthy. The injection of smad7 mRNA did not cause fragmentation of the cells (data not shown), but the embryos exhibited a decreased number of isl1-promoter-active cells in the TgBAC(isl1:GFP);Tg(myl7:Nls-mCherry) assay (Figure 6E,F and Figure 6—source data 1). Collectively, these results suggest that Bmp-dependent signaling is required for CPC fate determination.

Finally, to confirm the necessity of Bmp-Smad-regulated signaling during SHF formation, we treated the TgBAC(hand2:GFP);Tg(myl7:Nls-mCherry) embryos and the TgBAC(isl1:GFP);Tg(myl7:Nls-mCherry) embryos with a Bmp inhibitor, DMH1, from 14 hpf to 26 hpf. The efficiency of DMH1 was confirmed by decreased phosphorylation of Smad1/5/9 (Figure 6—figure supplement 2C). The expression of Isl1 and the promoter activity of hand2 were greatly reduced in the embryos treated with DMH1 (Figure 6—figure supplement 3C). The number of isl1-promoter-active SHF cells was decreased in the embryos treated with DMH1, whereas the number of isl1-promoter-inactive CMs in the DMH1-treated embryos was comparable to that in the control embryos (Figure 6G,H, and Figure 6—source data 1). We thus conclude that Lats1/2 restrict Yap1/Wwtr1-promoted Bmp2b-dependent signaling, which is required for both hand2- and isl1-promoter activity during SHF formation.

Discussion

Here, we show for the first time that the Hippo signaling pathway is involved in the determination of LPM cell fate by promoting venous pole identity and increases in atrial CM number (Figure 7). We show that Yap1/Wwtr1-promoted signaling increases the size of the SHF domain and that Lats1/2, by inhibiting Yap1/Wwtr1 activity, restrict it. We propose that the increased number of SHF cells in the lats1/2 DKO embryos may result from a change in fate determination of hand2-negative cells, which become hand2-positive cells in the boundary between ALPM and PLPM. Indeed, we found that the expression of the marker of blood-cell progenitors tal1 was repressed in the rostral region of PLPM in lats1/2 DKO embryos. lats1/2 mutants exhibited a subtle increase in the number of Isl1-positive atrial SHF cells, with no defect apparent in other organs. Importantly, despite the known role of Hippo signaling in CM proliferation during heart maturation, Hippo signaling was not found to affect cell proliferation during these early stages. Therefore, Hippo signaling contributes specifically to the determination of LPM differentiation.

A schematic illustration of Hippo signaling in the ALPM and the border of the ALPM and PLPM in wildtype (WT) and Lats1/2 double knockout (DKO) embryos.

In WT embryos, at the caudal part of the ALPM, Hippo signaling is inactive, whereas Tead-dependent transcription co-activated by Yap1/Wwtr1 is active and promotes bmp2b expression and hand2 expression. Cells expressing hand2 become the Isl1-positive SHF cells in the venous pole of the heart tube, and eventually populate the inflow tract. In the rostral region of the PLPM, Hippo signaling is active and hand2 expression is suppressed. Cells from this region do not become cells of the venous pole. In the Lats1/2 DKO, Hippo signaling is absent in the caudal part of the ALPM, and hand2 expression is increased. In the rostral region of the PLPM, Hippo signaling is absent and hand2 is expressed. hand2 expression in these cells promotes SHF specification, and these cells are integrated into the venous pole of the heart tube, and eventually populate the atrium of the heart.

https://doi.org/10.7554/eLife.29106.025

We speculate that Hippo signaling cooperates with other signaling pathways to determine HF formation. For example, hoxb5b expression in the forelimb field limits the extent of the HF at the posterior border of the ALPM, cells of which differentiate into atrial but not ventricular CMs (Waxman et al., 2008). Furthermore, other signals generated in the pronephric field in the intermediate mesoderm and in the angiogenic field in the rostral region of PLPM are important for the regulation of cell fate at the posterior HF boundary (Kimmel et al., 1990; Mudumana et al., 2008).

Our results also help to clarify the origin of venous pole CMs in zebrafish and its link to the SHF. In the mouse embryo, the anterior and posterior SHF cells differentiate into the OFT/right ventricular myocardium and the IFT/atrial myocardium, respectively (Galli et al., 2008; Verzi et al., 2005). The posterior-SHF in the HF is located caudally (Abu-Issa and Kirby, 2008; Galli et al., 2008). Tead-reporter activation occurred in the caudal zone of the ALPM, and these cells were shown to become Isl1-positive, hand2-promoter-active CMs that are localized to the venous pole of the atrium. Both sides of the HF located caudally contributed to the venous pole of the cardiac tube. Further, mammalian SHF cells have multi-potential to differentiate into endocardial cells and smooth muscle cells in addition to myocardial cells (Chen et al., 2009b). By generating BAC transgenic fish, we found that isl1-promoter-active cells were detected in the atrial myocardium, endocardium, and epicardium, but not in the ventricular myocardium. Together with previous reports, our results suggest that the properties of zebrafish Isl1-positive SHF cells are similar to those of mammalian posterior-SHF cells.

Lats1/2-Yap1/Wwtr1-Tead signaling regulates Bmp2b expression, which is necessary for the formation of hand2-promoter-active cells in the ALPM and Isl1-expressing cells. Although previous reports have shown that isl1-positive and mef2-positive cells reside at the venous pole (de Pater et al., 2009; Hinits et al., 2012), the molecular mechanism explaining how ALPM cells give rise to these CPCs at the venous pole has remained unclear. To date, a number of signaling molecules, such as TGFβ, FGF, and BMP, have been reported to regulate arterial pole formation in the zebrafish heart tube (de Pater et al., 2009; Hami et al., 2011; Zhou et al., 2011). We found that the Tead activation signal overlaps with the Bmp-reporter-positive signal in the ALPM. By analyzing lats1/2 mutants and yap1/wwtr1 mutants, we show that Hippo signaling controls bmp2b expression in the ALPM. Bmp-Smad inhibition expands the tal1 expression domain to restrict LPM fate (Gupta et al., 2006). Interestingly, hand2 expression is diminished in the alk3 mutant, which is affected in a Bmp type I receptor 1a, at 12 ss (de Pater et al., 2012). In our experiments, Bmp-Smad inhibition results in the suppression of hand2-promoter-activated GFP expression at 15 hpf. Therefore, we conclude that bmp2b expression is positively regulated by Yap1/Wwtr1, which balances the cell fate between the HF and the blood cell field at the boundary between the ALPM and the PLPM.

We believe that the downstream targets of the Hippo-dependent Bmp-mediated signal, especially the transcription-factor-(Smads)-dependent signal, might promote distinct functions that depend on their time of action. It has been shown previously that Bmp signaling is upstream of both hand2 and nkx2.5 in zebrafish, and that the expression of both mRNAs is lost in Bmp signaling mutants (de Pater et al., 2012; Kishimoto et al., 1997; Reiter et al., 2001). However, we found here that yap1/wwtr1 double mutants still have hand2 expression (although this is reduced) and normal nkx2.5 expression, even though Bmp signaling is decreased. Consistent with this possibility, Bmp signal-dependent dorso-ventral axis formation is not dependent on Lats1/2-Yap1/Wwtr1-Bmp signaling, suggesting that additional regulators of Bmp signaling are active in the ALPM. Another possibility is that Hippo signaling acts independently on both bmp2b expression and hand2 expression.

In summary, we demonstrate that the Yap1/Wwtr1-Tead signal promotes Bmp2b expression and that Smad-dependent signaling subsequently defines the ALPM (Figure 7). Because Hippo signaling restricts the border between the ALPM and the PLPM, Hippo signaling may account for the restriction of SHF formation and the subsequent SHF-derived Isl1-positive IFT atrial CMs.

Materials and methods

Key resources table
Reagent type (species)
or resource
DesignationSource or referenceIdentifiersAdditional information
Gene (Danio rerio)yap1NAZDB-GENE-030131–9710PMID:25313964
Gene (Danio rerio)wwtr1NAZDB-GENE-051101–1PMID:28350986
Gene (Danio rerio)lats1NAZDB-GENE-050523–2PMID:19842174
Gene (Danio rerio)lats2NAZDB-GENE-050119–6PMID:19842174
Gene (Danio rerio)smad7NAZDB-GENE-030128–3
Gene (Danio rerio)bmp2bNAZDB-GENE-980526–474PMID:19000838
Gene (Danio rerio)bmp4NAZDB-GENE-980528–2059
Gene (Danio rerio)gata4NAZDB-GENE-980526–476
Gene (Danio rerio)nkx2.5NAZDB-GENE-980526–321PMID:23444361
Gene (Danio rerio)etv2NAZDB-GENE-050622–14
Gene (Danio rerio)tal1NAZDB-GENE-980526–501
Gene (Danio rerio)isl1NAZDB-GENE-980526–112PMID:19395641
Gene (Danio rerio)hoxb5bNAZDB-GENE-000823–6PMID:19081079
Strain, strain background
(Danio rerio)
ABZIRCZDB-GENO-960809–7;
RRID:ZIRC_ZL1
Genetic reagent
(Danio rerio)
hand2 BAC:GFPPMID:20627079ZDB-ALT-110128–40;
RRID:ZFIN_ZDB-ALT-110128-40
Genetic reagent
(Danio rerio)
hand2 BAC:tdEosFPThis paperBacPac Resources:CH211-
95C16 BAC
Genetic reagent
(Danio rerio)
isl1 BAC:GFPThis paperBacPac Resources:CH211-
219F7 BAC
Genetic reagent
(Danio rerio)
eef1a1l1:galdb-hTEAD2ΔN-
2A-mCherry
PMID:25313964ZDB-FISH-150901–27167
Genetic reagent
(Danio rerio)
myl7:galdb-hTEAD2ΔN-2A-
mCherry
This paper
Genetic reagent
(Danio rerio)
uas:GFPPMID:18202183
Genetic reagent
(Danio rerio)
uas:mRFP1PMID:18202183
Genetic reagent
(Danio rerio)
myl7:Nls-mCherryPMID:25313964ZDB-GENO-150218–2
Genetic reagent
(Danio rerio)
myh6;Nls-tdEosFPThis paper
Genetic reagent
(Danio rerio)
myl7:EGFPThis paper
Genetic reagent
(Danio rerio)
BRE:GFPPMID:21337469ZDB-ALT-110308–1;
RRID:ZFIN_ZDB-ALT-110308-1
Genetic reagent
(Danio rerio)
tol2 transposase mRNAPMID:1523996150 pg injection
Genetic reagent
(Danio rerio)
ytip-mCherry mRNAPMID:25313964200 pg injection
Genetic reagent
(Danio rerio)
lats1-atg MOPMID:19842174,
GeneTools
ZDB-MRPHLNO-100415–21.2 ng injection
Genetic reagent
(Danio rerio)
lats2-atg MOPMID:19842174,
GeneTools
ZDB-MRPHLNO-100415–41.2 ng injection
Genetic reagent
(Danio rerio)
ajuba-atg MOPMID:22771034,
GeneTools
ZDB-MRPHLNO-120821–58 ng injection
Genetic reagent
(Danio rerio)
control MOStandard control
oligo, GeneTools
5 ng injection
AntibodyAnti-GFP antibodyAbcamAbcam:ab13970;
RRID:AB_300798
(1:300)
AntibodyAnti-mCherry antibodyClontechClontech:632543;
RRID:AB_2307319
(1:300)
AntibodyAnti-Isl1 antibodyGenetexGenetex:GTX128201(1:100)
AntibodyAnti-pSmad1/5/9
antibody
Cell Signaling
Technology
Cell Signaling Technology:
13820S
(1:100)
AntibodyAnti-chicken Alexa Fluor
488 IgG
Thermo Fisher
Scientific
Thermo Fisher Scientific:
A-11039; RRID:AB_142924
(1:300)
AntibodyAnti-mouse Alexa Fluor
546 IgG
Thermo Fisher
Scientific
Thermo Fisher Scientific:
A-11030; RRID:AB_2534089
(1:300)
AntibodyAnti-rabbit Alexa Fluor
633 IgG
Thermo Fisher
Scientific
Thermo Fisher Scientific:
A-21070; RRID:AB_2535731
(1:300)
Recombinant DNA
reagent
pCR4-bluntTOPOThermo Fisher
Scientific
Thermo Fisher Scientific:
K287520
Recombinant DNA
reagent
pTol2PMID:18202183
Recombinant DNA
reagent
pcDNA3.1Thermo Fisher
Scientific
Thermo Fisher Scientific:
V790-20
Recombinant DNA
reagent
pRedETGene Bridges
Recombinant DNA
reagent
pCS2ClontechRRID:SCR_007237
Commercial assay
or kit
mMessage mMACHINE
kit
Thermo Fisher
Scientific
Thermo Fisher Scientific:
AM1340
Commercial assay
or kit
KOD FX Neo DNA
polymerase
TOYOBOTOYOBO:KFX-201
Commercial assay
or kit
Click-iT EdU Alexa 647
Imaging Kit
Thermo Fisher
Scientific
Thermo Fisher Scientific:
C10340
Commercial assay
or kit
DIG RNA labeling litRocheSigma-Aldrich:11175025910
Commercial assay
or kit
BM-purpleRocheSigma-Aldrich:11442074001
Commercial assay
or kit
QuantiFast SYBR Green
PCR kit
QiagenQiagen:204054
Chemical compound,
drug
DMH1CalbiochemSigma-Aldrich:20364610 μM addition
Chemical compound,
drug
PTUSigma-AldrichSigma-Aldrich:P7629
Software, algorithmGraphPad Prism 7GraphPad SoftwareRRID:SCR_002798
Software, algorithmImaris ver.8.4.1BitplaneRRID:SCR_007370
OtherMultiNA microchip
electrophoresis system
SHIMADZU

Zebrafish (danio rerio) strains, transgenic lines, and mutant lines

The experiments using zebrafish were approved by the institutional animal committee of National Cerebral and Cardiovascular Center (permit number: 17003) and performed according to the guidelines of the Institute. We used the AB strain as wildtype. The following zebrafish transgenic lines were used for experiments: Tg(eef1a1l1:galdb-hTEAD2ΔN-2A-mCherry) fish (Fukui et al., 2014), Tg(myl7:Nls-mCherry) fish (Fukui et al., 2014), TgBAC(hand2:GFP) fish (Yin et al., 2010), Tg(BRE:GFP) fish (Collery and Link, 2011), Tg(uas:mRFP1) fish (Asakawa et al., 2008), and Tg(uas:GFP) fish (Asakawa et al., 2008). The Tg(myl7:galdb-hTEAD2ΔN-2A-mCherry) fish, Tg(myh6:Nls-tdEosFP) fish, Tg (myl7:EGFP) fish, and TgBAC(isl1:GFP) fish were generated as described in the experimental procedures. The knockout alleles ncv107 for lats1, ncv108 for lats2, and ncv117 for yap1 genes were generated by TALEN techniques as described in the experimental procedures. The ncv114 allele for wwtr1 was previously described by Nakajima et al (2017).

Image acquisition and image processing

To obtain the images of embryos, the pigmentation of the embryos was suppressed by the addition of 1-phenyl-2-thiourea (PTU) (Sigma-Aldrich, St. Louis, MO) into breeding E3 media. Embryos were dechorionated and mounted in 1% low-melting agarose dissolved in E3 medium. Confocal images of 2.0 μm steps were taken with a FV1200 confocal microscope system (Olympus, Tokyo, Japan) equipped with a water immersion 20x lens (XLUMPlanFL, 1.0 NA, Olympus). Images were processed with a FV10-ASW 4.2 viewer (Olympus). The distance between the hand2-positive region of the ALPM and the PLPM was measured using DP2-BSW software (Olympus). Cell-tracking data containing nuclei positions were analyzed using Imaris8.4.1 software (Bitplane, Zurich, Switzerland).

Generation of knockout zebrafish by TALEN

To develop knockout zebrafish, we used transcription activator-like effector nuclease (TALEN) Targeter 2.0 (https://tale-nt.cac.cornell.edu) to design a TALEN pair that targets lats1, lats2, and yap1. The target sequence of TAL-lats1, TAL-lats2, and TAL-yap1 were 5′-TCAGCAAATGCTGCAGGAGATccgagagagcctgcgaAACCTCTCCCCGTCCTCCAA-3′, 5′-TCTCGAGGAGAGGGTGgtcgaggtggagactCAAAGGGCAAAGACCA-3′, and 5′-CCGAACCAGCACAACCctccagccggccaccagaTCGTCCATGTTCGGGG-3′, respectively (capital letters are the sequences of the left [TAL-lats1-F, lats2-F, and yap1-F] and right [TAL-lats1-R, lats2-R, and yap1-R] arms, respectively). These expression plasmids of the TALEN-pair were constructed by pT3TS-GoldyTALEN. TALEN mRNAs were synthesized in vitro using a T3 mMessage mMACHINE kit (Thermo Fisher Scientific, Waltham, MA). To induce double-strand breaks in the target sequence, 50 pg of TAL-lats1-F / -lats1-R mRNAs, TAL-lats2-F / -lats2-R mRNAs, or TAL-yap1-F / -yap1-R mRNAs were injected into one- to two-cell stage transgenic embryos. Each injected founder (F0) fish was outcrossed to wildtype fish to obtain the F1 progeny from individual founders. The generation of wwtr1 knockout zebrafish has been reported previously (Nakajima et al., 2017). To analyse TALEN-induced mutations, genomic DNA from F1 embryos was lysed by 50 μl of NaOH solution (50 mM) at 95°C for 5 min, and 5 μl of Tris-HCl (pH8.0, 1.0 M) was added on ice for 10 min. After centrifugation (13,500 rpm, 5 min), PCR reaction was performed using KOD FX Neo DNA polymerase (TOYOBO, Osaka, Japan). The genotyping PCR primers were used for amplification: lats1 (5′-GGCACTTAACATATGCTTTTACATG-3’ and 5′-TTTGCTGCTGTCTGCGGAGCTGTT-3′); lats2 (5′-AGAGTTTGTGTGAGAGAAAACAGG-3’ and 5′-GCATTGACCAGATCCTGTAGCATC-3′); yap1 (5′-TCCTTCGCAAGGCTTGGATAATTG-3’ and 5′-TTGTCTGGAGTGGGACTTTGGCTC-3′); wwtr1 (5′-GGACGAAAAACAGGAAAAGTTC-3’ and 5′-ACTGCGGCATATCCTTGTTC-3′). These amplified PCR products were analyzed using a MCE-202 MultiNA microchip electrophoresis system (SHIMADZU, Kyoto, Japan) with the DNA-500 reagent kit (SHIMADZU).

Microinjection of oligonucleotide and mRNA

We injected 200 pg ytip-mCherry mRNA (Fukui et al., 2014), 100 or 200 pg zebrafish-smad7 mRNA, 1.2 ng lats1-atg MO (5′-CCTCGGGTTTCTCGGCCCTCCTCAT-3′) (Chen et al., 2009a), 1.2 ng lats2-atg MO (5′-CATGAGTGAACTTGGCCTGTTTTCT-3′) (Chen et al., 2009a), 8 ng ajuba-atg MO (5′-TGAGTTTGATGCCAAGTCGATCCAT-3′) (Witzel et al., 2012), and 5 ng control MO (5′-CCTCTTACCTCAGTTACAATTTATA-3′) as previously reported (Fukui et al., 2014). These morpholinos were purchased from Gene Tools (Philomath, OR). Capped mRNAs were synthesized using the SP6 mMessage mMachine system (Thermo Fisher Scientific). Microinjection was performed using FemtoJet (Eppendorf, Hamburg, Germany). MOs, mRNA, and Tol2 plasmids were injected into blastomeres at the one- to two-cell stage.

EdU incorporation assay

The TgBAC(isl1:GFP);Tg(myl7:Nls-mCherry) embryos injected with control MO or lats1/2 MOs were incubated with 2 mM of 5-ethynyl-2-deoxyuridine (EdU) from 14 to 26 hpf or from 20 to 36 hpf, and subsequently fixed using 4% PFA at 96 hpf. EdU-incorporated cells were labelled by Click-iT EdU Alexa Fluor 647 Imaging Kits (Thermo Fisher Scientific) following the manufacturer’s instructions. Images were taken using the FV1200 confocal microscope system. The number of EdU-positive isl1-promoter-active CMs was determined by counting the number of cells with overlapping Alexa Fluor 647-positive signal, isl1-promoter-activated signal and myl7-promoter-activated signal.

Whole-mount in situ hybridization (WISH)

The antisense hand2, isl1, bmp2b, bmp4, gata4, nkx2.5, etv2, tal1, and hoxb5b RNA probes labeled with digoxigenin (DIG) were prepared using an RNA labeling kit (Roche, Basel, Switzerland). WISH was performed as previously described (Fukui et al., 2014). Colorimetric reaction was carried out using BM purple (Roche) as the substrate. To stop the reaction, embryos were washed with PBS-T, fixed with 4% PFA for 20 min at room temperature, and subsequently immersed in glycerol. Images were taken using a SZX-16 Stereo Microscope (Olympus).

Immunohistochemistry

Embryos at 14 hpf and 26 hpf were fixed by MEMFA (3.7% formaldehyde, 0.1 M MOPS, 2 mM EGTA, 1 mM MgSO4) for 2 hr at room temperature. After fixation, the solution was changed to 50% methanol/MEMFA for 10 min, then changed to 100% methanol at room temperature, and then stored in 100% methanol at –30°C overnight. After rehydration, embryos were washed three times for 10 min in PBBT (PBS with 2 mg/mL BSA and 0.1% TritonX-100). Embryos were blocked in PBBT with 10% goat serum for 60 min at room temperature, and subsequently incubated overnight at 4°C with primary antibodies, 1:300 diluted chicken anti-GFP antibody (ab13970, Abcam, Cambridge, UK), 1:300 diluted mouse anti-mCherry antibody (632543, Clontech, Mountain View, CA), and 1:100 diluted rabbit anti-Islet1 antibody (GTX128201, Genetex, Irvine, CA) or 1:100 diluted rabbit anti-pSmad1/5/9 antibody (13820S, Cell Signaling TECHNOLOGY, Danvers, MA) in blocking solution. Embryos were washed with PBBT five times over the course of 2 hr, with blocking solution for 60 min at room temperature, and incubated overnight at 4°C with secondary antibodies, anti-chicken Alexa Fluor 488 IgG (A-11039, Thermo Fisher Scientific), anti-mouse Alexa Fluor 546 IgG (A-11030, Thermo Fisher Scientific), and anti-rabbit Alexa Fluor 633 IgG (A-21070, Thermo Fisher Scientific) diluted 1:300 in blocking solution. Embryos were washed with PBBT five times over the course of 2 hr and stored in PBS at 4°C prior to imaging.

Quantitative real-time PCR (q-PCR)

Total RNAs were collected from whole-embryonic cells using TRizol (Thermo Fisher Scientific) following the manufacturer’s instructions. For q-PCR, reverse transcription and RT-PCR were performed with the QuantiFast SYBR Green PCR kit (Qiagen, Hilden, Germany) in the Mastercycler Realplex (Eppendorf). The following primer set was used for amplification: nkx2.5-S (5′-GCTTTTACGCGAAGAACTTCC-3′), nkx2.5-AS (5′-GATCTTCACCTGTGTGGAGG-3′); gata4-S (5′-AAGGTCATCCCGGTAAGCTC-3′), gata4-AS (5′-TGTCACGTACACCGGAGAAG-3′); hand2-S (5′-TACCATGGCACCTTCGTACA-3′), hand2-AS (5′-CCTTTCTTCTTTGGCGTCTG-3′); eef1a1l1-S (5′-CTGGAGGCCAGCTCAAACAT-3′), eef1a1l1-AS (5′-ATCAAGAAGAGTAGTACCGCTAGCATTAC-3′) (Fukui et al., 2014).

Plasmids cDNA fragments encoding zebrafish Hand2, Isl1, Bmp2b, Bmp4, Gata4, Nkx2.5, Etv2, Tal1, Hoxb5b and Smad7 were amplified by PCR using a cDNA library derived from zebrafish embryos and subcloned into a pCR4 Blunt TOPO vector (Thermo Fisher Scientific). The following primer set were used for amplification: hand2-S (5′-CGGGATCCCGCCATGAGTTTAGTTGGAGGGTT-3’ [containing BamHI sequence]), hand2-AS (5′-GCTTTAGTCTCATTGCTTCAGTTCC-3′); isl1-S (5′-GCTCTAGACCTTACTTTCTTGACATGGGAGAC-3’ [containing XbaI sequence]), isl1-AS (5′-GGACTGGTCGCCACCATTGGAGTA-3′); bmp2b-S (5′-ATGTCGACACCATGGTCGCCGTGGTCCGCGCTCTC-3’ [containing SalI sequence]), bmp2b-AS (5′-TCATCGGCACCCACAGCCCTCCACC-3′); bmp4-S (5′-CGGGATCCCATGATTCCTGGTAATCGAATGC-3’ [containing BamHI sequence]), bmp4-AS (5′-CATTTGTACAACCTCCACAGCAAG-3′); gata4-S (5′-GTGAATTCATGTATCAAGGTGTAACGATGGCC-3’ [containing EcoRI sequence]), gata4-AS (5′-GAGCTTCATGTAGAGTCCACATGC-3′); nkx2.5-S (5′-GCTCTAGATTCCATGGCAATGTTCTCTAGCCAA-3’ [containing XbaI sequence]), nkx2.5-AS (5′-GATGAATGCTGTCGGTAAATGTAG-3′); etv2-S (5′-GTGAATTCCTGGATTTTACACAGAAGACTTCAGA-3’ [containing EcoRI sequence]), etv2-AS (5′-CCACGACTGAGCTTCTCATAGTTC-3′); tal1-S (5′-GTGAATTCGAAATCCGAGCAATTTCCGCTGAG-3’ [containing EcoRI sequence]), tal1-AS (5′-CTTAGCATCTCCTGAAGGAGGTCGT-3′); hoxb5b-S (5′-GTGAATTCCCAAATGAGCTCTTATTTTCTAAACTCG-3’ [containing EcoRI sequence]), hoxb5b-AS (5′-GATGTGATTTGATCAATTTTGAAACGCGC-3′); smad7-S (5′-AGGGATCCTCCCGCATGTTCAGGACCAAACGAT-3’ [containing BamHI sequence]), smad7-AS (5′-GAAGGCCTTTATCGGTTATTAAATATGACCTCTAACC-3’ [containing StuI sequence]). The cDNA of zYtip was previously amplified and cloned into the pCS2 vector (Clontech) (Fukui et al., 2014). The DNA encoding Smad7 was subcloned into the pCS2 vector to construct the pCS2-smad7. All of the cDNAs amplified by PCR using cDNA libraries were sequenced. Mutations were also confirmed by sequencing.

Generation of transgenic lines

To monitor atrial CM development, we established a transgenic (Tg) zebrafish line expressing a nuclear localization signal (Nls)-tagged tandem Eos fluorescent protein under the control of the myosin heavy chain 6 (myh6) promoter; Tg(myh6:Nls-tdEosFP). pTol2-myh6 vector was constructed by modifying the pTol2 vector and inserting the myh6 promoter as a driver of the expression of the target molecule. The primers used to amplify the myh6 promoter were 5′-AGAGCTAAAGTGGCAGTGTGCCGAT-3’ and 5′-TCCCGAACTCTGCCATTAAAGCATCAC-3′. An oligonucleotide encoding Nls derived from SV40 (PKKKRKV) was inserted into pcDNA-tdEosFP (MoBiTec, Göttingen, Germany) to generate the plasmids expressing Nls-tagged tandem Eos fluorescent protein (Nls-tdEosFP). The Nls-tdEosFP cDNA was subcloned into the pTol2-myh6 vector to construct the pTol2-myh6:Nls-tdEosFP plasmids.

To monitor CM-specific Yap1/Wwtr1-dependent transcriptional activation, we developed a transgenic fish that expresses human (h) TEAD2 lacking the amino-terminus (1–113 aa) and fused with the Gal4 DNA-binding domain followed by 2A mCherry under the control of the myosin light polypeptide 7 (myl7) promoter; Tg(myl7:galdb-hTEAD2ΔN-2A-mCherry). This Tg fish was crossed with Tg(uas:GFP) reporter fish to obtain Tg(myl7:galdb-hTEAD2ΔN-2A-mCherry);Tg(uas:GFP). The pTol2-myl7 vector and the pcDNA3.1 vector containing human TEAD2ΔN cDNA fused to the DNA-binding domain of Gal4 (pcDNA3.1-galdb-hTEAD2ΔN) were constructed as previously described (Fukui et al., 2014). The Gal4db-hTEAD2ΔN cDNA was subcloned into the pTol2-myl7 vector to construct the pTol2-myl7:galdb-hTEAD2ΔN plasmids.

To monitor CM development, we developed a transgenic line that expresses EGFP under the control of the myl7 promoter; Tg(myl7:EGFP). The EGFP was subcloned into a pTol2-myl7 vector to construct the pTol2-myl7:EGFP plasmids. All of the cDNAs amplified by PCR using cDNA libraries were confirmed by DNA sequencing.

To monitor SHF development, we established a transgenic line that expressed GFP under the control of isl1 BAC promoter/enhancer; the TgBAC(isl1:GFP). pRedET plasmid (Gene Bridges, Heidelberg, Germany) was introduced into E. coli containing a CH211-219F7 BAC clone encoding the isl1 gene (BacPAC resources) by electroporation (1800V, 25 mF, 200 Ω) to increase the efficiency of homologous recombination, as previously described (Ando et al., 2016). Tol2 long terminal repeats in opposite directions flanking an ampicillin resistance cassette were amplified by PCR using Tol2_amp as a template, and these sequences were inserted into the BAC vector backbone. The cDNA encoding both GFP and a kanamycin resistance cassette (GFP_KanR) was amplified by PCR using a pCS2-GFP_KanR plasmid as a template, and inserted into the start ATG of the isl1 gene. Primers to amplify the GFP_KanR for isl1 gene were 5′-gggccttctgtccggttttaaaagtggacctaacaccgccttactttcttACCATGGTGAGCAAGGGCGAGGAG-3’ and 5′-aaataaacaataaagcttaacttacttttcggtggatcccccatgtctccTCAGAAGAACTCGTCAAGAAGGCG-3’ (small letters are the homology arm to the BAC vector, whereas capital letters are the primer binding site to the template plasmid).

Tol2-mediated zebrafish transgenesis was performed by injecting 30 pg of the transgene plasmid together with 50 pg tol2 transposase mRNA, followed by subsequent screening of F1 founders and establishment of single-insertion transgenic strains through selection in F3 generations.

Photoconversion

We performed photoconversion experiments by examining the transient hand2-promoter-dependent expression of tdEosFP. Tg(myl7:EGFP) embryos were injected with 30 pg of pTol2-hand2 BAC:tdEosFP plasmid along with 50 pg tol2 transposase mRNA. To trace ALPM cells, the caudal region of the left or right ALPM was photoconverted by a 405 nm laser at 15 hpf (12 ss). The photoconverted cells expressing red fluorescence were traced in the heart region at 52 hpf.

To construct the pTol2-hand2 BAC:tdEosFP, pRedET plasmid was introduced into E. coli containing a CH211-95C16 BAC clone encoding the hand2 gene (BacPAC resources) by electroporation. Tol2 long terminal repeats in opposite directions flanking an ampicillin resistance cassette were amplified by PCR using Tol2_amp as a template, and were inserted into the BAC vector backbone. The cDNA encoding tdEosFP together with a kanamycin resistance cassette (tdEosFP_KanR) was amplified by PCR using the pCS2-tdEosFP_KanR plasmid as a template, and then inserted into the start ATG of the hand2 gene. Primers to amplify the tdEosFP_KanR for the hand2 gene were 5′-ccaaagcgtactccgtctgtggttcgccgtagggtatagacaagtctgtcACCATGAAGATCAACCTCCGTATGGAAG-3’ and 5′-tagccgtcatggtgcatcacagggtggtggggaaaccctccaactaaactTCAGAAGAACTCGTCAAGAAGGCG-3’ (small letters represent the homology arm to BAC vector, and capital letters the primer binding site to the template plasmid).

Chemical treatment

The TgBAC(hand2:GFP);Tg(myl7:Nls-mCherry) embryos and the TgBAC(isl1:GFP);Tg(myl7:Nls-mCherry) embryos were treated with 10 μM DMH1 (203646, Sigma-Aldrich), an inhibitor of Bmp signaling, from 14 hpf to 26 hpf. As a control, embryos were incubated in E3 solution containing DMSO. These embryos were imaged at 26 hpf.

Data analysis and statistics

Data were analyzed using GraphPad Prism 7 (GraphPad Software, La Jolla, CA). All columns shown in histograms represent a mean ± SEM. The statistical significance of multiple groups was determined by one-way ANOVA with Bonferroni’s post hoc test. The numbers of atrial and ventricular CMs at 74 hpf were analyzed by Student’s t-test. The statistical significance of two groups was determined by Student’s t-test.

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Decision letter

  1. Deborah Yelon
    Reviewing Editor; University of California, San Diego, United States

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "Hippo signaling restricts cells in the second heart field that differentiate into Islet-1-positive atrial cardiomyocytes" for consideration by eLife. Your article has been reviewed by three peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Didier Stainier as the Senior Editor. The reviewers have opted to remain anonymous.

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

Summary:

In this manuscript, Fukui et al. examine the role of the Hippo signaling pathway in regulating development of the inflow tract (IFT) in the zebrafish heart. Using a series of elegant experiments and detailed quantification, the authors demonstrate that Hippo signaling is active in the developing IFT and that enhanced Hippo signaling results in an increased number of IFT cells, as labelled by hand2 and islet1 expression, as well as increased atrial cell number at later stages. The mechanism for the increase in cell number is not through proliferation but rather through increased differentiation of cardiomyocytes. Furthermore, the authors demonstrate that Bmp signaling is increased in the context of enhanced Hippo signaling, and they propose to place Bmp signaling downstream of Hippo and upstream of hand2 and islet1. Previous work has shown that Hippo signaling in mouse embryos determines heart size by regulating cardiomyocyte proliferation. In contrast, this work suggests that Hippo can also affect heart size by regulating specification of IFT cells. Thus, this study is significant on several levels: it expands our understanding of the roles of the Hippo signaling pathway in regulating heart size, it adds to our understanding of the pathways that regulate the formation of the IFT, and it has the potential to provide broader insights into the mechanisms that control of cardiac differentiation and patterning. However, certain elements of the authors' conclusions are not fully supported by the data presented here. Several issues require further investigation and/or clarification in order to make a clear case for their model.

Essential revisions:

1) Importantly, the authors need to provide further evidence to support their conclusions regarding the differential fates of the left and right caudal ALPM. Previous cell tracking work has shown that cells from the left LPM form the dorsal side of the cardiac tube while cells from the right LPM form the ventral side of the cardiac tube (PMID: 18267096). Different from the description in the current manuscript, the older work showed that both the right and left LPM cells contributed to the venous pole. However, the dorsal side of the tube extends more anteriorly compared to the ventral side. Looking closely at the data provided by the authors (e.g. by playing the last 3 hours (equivalent of 2 secs) of their movie backward and forward rapidly), one gets the impression that the right caudal ALPM also contributes to the venous pole at the ventral side. Likely due to the tissue depth at the ventral surface, the fluorescence intensity of these cells is weaker and, therefore, difficult to track. To definitively state that the left and right sides contribute to the venous and arterial poles, respectively, the authors should provide higher resolution (and 3D) tracking of both the right and left ALPM cells. In addition, complementary lineage tracing experiments (using photoconversion or other cell labeling techniques) should be employed to follow the cells labeled in the right and left ALPM to their destination in the heart at 48 hpf.

2) The authors highlight the expression of the Tead reporter in the left caudal ALPM, but it is not clear whether or not this reporter is expressed in the right caudal ALPM. This needs to be clarified.

3) The ectopic hand2 expression and bmp2 expression in lats1/2 mutants is bilateral and not restricted to the left ALPM (Figures 5 and 6). How does this reconcile with the authors' suggested laterality of Hippo signaling (as shown in their model in Figure 7)?

4) In Figure 6, the authors show enhanced expression of bmp2b in lats1/2 mutants and loss of bmp2 expression in yap/wwtr mutants. The effects are visible in the entire ALPM and head region, while their data shown in Figure 3 suggests that Yap1 is only active in the posterior ALPM. This discrepancy is not addressed. How do the authors fit these data with their model in which Hippo signaling activity is different in the posterior versus anterior ALPM?

5) It's possible that an increase in signaling through either pathway (Hippo or Bmp) would result in increased cells at the IFT. Is there increased TEAD reporter activity following bmp2b heat-shock?

6) To reduce BMP signaling, the authors inject mRNA for smad7 (inhibitory Smad). It is not clear why the authors try to block BMP signalling during early stages of development since BMP signaling is required for proper dorsal-ventral patterning. Did the authors observe DV patterning defects in these experiments?

7) The authors imply that lats1 and lats2 have a special influence on the Isl1-positive atrial cells and do not affect the numbers of the Isl1-negative atrial cells. However, this is not clearly demonstrated by the data presented here: the total number of atrial cells is presented at one stage, and the number of Isl1-positive cells is presented at a substantially earlier stage. Does the change in the number of Isl1-positive cells account for the entirety of the difference in atrial size? Similarly, is the impact of overexpressing bmp2b and/or adding DMH1 specific to the Isl1-positive population, or do these manipulations affect the entire atrium?

8) The role of Yap1/Wwtr1 in promoting hand2 expression needs further clarification. The authors point to the reduction of hand2 expression in yap1;wwtr1 mutants, but it is unclear how specifically this phenotype relates to the regulation of hand2. Is the entire ALPM reduced in yap1;wwtr1 mutants? In addition, the relationship between the cardia bifida phenotype and the reduced hand2 expression in the yap;wwtr1 mutant should be more clearly articulated. The yap;wwtr1 mutant appears smaller than the control embryo shown. Corresponding bright field images would be helpful here.

[Editors' note: further revisions were requested prior to acceptance, as described below. The authors’ plan for revisions was approved and the authors made a formal revised submission.]

Thank you for sending your revised article entitled "The Hippo signaling suppresses the differentiation of cardiac precursor cells from lateral plate mesoderm prior to establishment of the heart field in zebrafish" for peer review at eLife. Your revised article has been evaluated by its original three peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation is being overseen by Didier Stainier as the Senior Editor.

The reviewers have discussed their reviews with one another, and their discussion has highlighted some remaining issues (elaborated below) that still need to be addressed. Since some of these revisions would likely require you to perform new experiments, we ask that you respond to this letter within the next two to three weeks with a specific action plan and timetable for the completion of the additional work. The Reviewing Editor and reviewers will then consider your plans, assess the feasibility of an effective response to their concerns, and offer a binding recommendation (such as an official invitation to submit a second revision of your manuscript to eLife). Alternatively, if you would prefer to submit your manuscript elsewhere, rather than revising it again for eLife, please let us know.

In this revised manuscript, the authors have made multiple amendments to their original submission, including new data and new interpretations, and many of these have helped to clarify issues from the original manuscript that needed to be addressed. However, several concerns remain that require further attention. Importantly, while the current version of the manuscript certainly demonstrates an influence of lats1 and lats2 on the development of the venous pole, it stops short of clearly articulating a mechanistic connection between the Hippo pathway and the differentiation of venous pole cells. Specifically, some of the data shown appear to be inconsistent with or to contradict the authors' model, as described below.

Essential revisions:

1) From the authors' data, it is clear that there is a positive relationship between Bmp signaling and the formation of venous pole cells. It is also clear that Hippo signaling has a negative influence on bmp2b expression. However, it is not yet clear whether the caudal boundary of hand2 expression in the ALPM is set by Hippo signaling repressing bmp2b expression at that location. While it may be challenging to test a model that relates Hippo signaling to bmp2b expression directly, it would be beneficial for the authors to clarify aspects of their story that are not consistent with that model. For example, while bmp2b expression is increased in lats1;2 mutants, it does not appear to be expanded, and this seems to contradict the authors' model. Can the authors clarify this, or adjust their model accordingly?

2) The authors conclude that Yap1 activity in the ALPM induces Bmp signaling and that Bmp signaling activates hand2 and Isl1 expression, but this conclusion is too simple. Published data in zebrafish have shown that Bmp signaling is upstream of both hand2 and nkx2.5. (Both are lost in Bmp signaling mutants.) Yap1/wwtr1 double mutants still have hand2 expression (although reduced) and normal nkx2.5 expression. The authors do not address whether Bmp signaling is affected in the Yap1/wwtr1 mutants using the bre:gfp reporter line (they only show that bmp2b expression is reduced in the ALPM). Neither do they try to rescue the Yap1/wwtr1 phenotype with Bmp overexpression. The proposed model suggesting that Yap1 signaling is upstream of Bmp signaling is therefore an oversimplification and needs adjustment.

3) The phenotype shown for an embryo overexpressing smad7 mRNA does not seem to be a standard BMP-induced dorsalization phenotype, which is surprising. The appearance of the embryo in Figure 6—figure supplement 3A and the fragmented cells in Figure 6H suggest that the overexpression of smad7 may be causing cell death. Additionally, the cardiac phenotype attained after smad7 overexpression was not quantified, weakening the conclusions made from this experiment. If the authors intend to demonstrate cell-autonomous epistasis, they will likely need to use another approach. (Perhaps overexpression of a dominant-negative Bmp receptor?)

4) In the experiment using DNA injection to induce mosaic Bmp2b expression (Figure 6—figure supplement 2A), the cells expressing bmp2b-2A-mCherry look fragmented, suggesting toxicity. The tissue on the left side also looks necrotic and the Yap1 reporter is very weak, while the authors state that it is unaffected. These results are not appropriate for inclusion in the manuscript. It is unclear why the authors did not use the available hsp70:bmp2b transgenic line to generate mosaic embryos through transplantation.

5) In their revised manuscript, the authors conclude that venous pole cells originate on both sides of the ALPM, but they also state that "more cells moved from the left ALPM to the venous pole compared to the cells coming from the right ALPM". It is not clear that this semi-quantitative comparison is warranted, since it does not seem as if the authors tracked all of the cells on both sides of the ALPM. In Video 1, the authors compare five cells on the left to two cells on the right and state that this is representative of six independent experiments. How do the numbers compare among the six experiments? Is this a quantitatively reproducible observation? Do the photoconversion experiments (Figure 3D) reinforce the view that more venous pole cells come from the left than from the right? It is challenging to make a quantitative conclusion from this type of mosaic analysis. The authors should clarify the basis for this quantitative comparison between the left and right sides, or, alternatively, adjust their interpretation. Additionally, it would be helpful if the authors could clarify what criteria they used to define whether or not a tracked cell (or a photoconverted cell) became part of the venous pole in these experiments. Do the seven tracked cells in Video 1 account for the entirety of the venous pole?

6) The authors have revised their title and Abstract to accompany the changes to their model that they have incorporated into their revised manuscript. It may be beneficial to consider revising these further in order to better emphasize the key points that are most strongly supported by the current text. For example, the title claims that Hippo signaling "suppresses the differentiation of cardiac precursor cells" but it is not clear that suppression of differentiation is shown here. More venous pole cells are produced when Hippo signaling is inhibited, but it is not evident whether this occurs through alteration of differentiation (as opposed to alteration of specification or proliferation). To make this claim, the authors would need to provide additional data in support of this conclusion. Also, the authors emphasize that the activity of Hippo signaling occurs "prior to establishment of the heart field", but it is not clear what they mean by this phrase. When do they consider the heart field to be established, and what is the significance of its establishment as it relates to the conclusions of their manuscript? Overall, it would be more effective if the title and Abstract emphasized formation of the venous pole specifically, as that seems to be the focus of the manuscript.

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

Thank you for resubmitting your work entitled "Hippo signaling determines the number of atrial cells that originate from the anterior lateral plate mesoderm in zebrafish" for further consideration at eLife. Your revised article has been favorably evaluated by Didier Stainier (Senior editor), and three reviewers, one of whom is a member of our Board of Reviewing Editors.

The manuscript has been improved through its second round of revision, both by adding new data and by removing less convincing elements. Altogether, the revisions have streamlined and strengthened the overall clarity of the message regarding the impact of the Hippo signaling pathway on the number of venous pole cells in zebrafish, while toning down some of the more speculative or inconsistent aspects. However, there are some remaining issues that need to be addressed, as outlined below:

1) With each revision of this manuscript, the authors have modified their title and Abstract to accompany the changes in their message. Further modification would help the current title to fit more closely with the main point of the paper, which revolves around the pathways regulating the number of venous pole (Isl1+) cells. The new title and Abstract seem to emphasize the whole atrium, even though the take-home message of the data is focused on the venous pole. Further adjustment is needed to align the title and Abstract with the rest of the manuscript.

2) The data that are provided to argue that overexpression of smad7 (as in Figure 6) is non-toxic are not convincing. The authors have undertaken control experiments, examining whether embryos injected with smad7 mRNA look healthy, but this context is quite different from the mosaic scenario presented in the experiments, in which three different mRNAs are co-injected. It remains unexplored whether this cocktail is causing cell death (and therefore loss of hand2 expression) and therefore no conclusion can be drawn from this experiment. Therefore, the experiments utilizing smad7 overexpression should be removed from the manuscript.

3) It appears premature for the authors to conclude a direct, linear relationship connecting Hippo signaling, Bmp signaling, and Hand2 expression. Without a formal epistasis experiment (e.g. DMH1 treatment in lats1/2 LOF and measuring the hand2 response), it does not seem appropriate to suggest this hierarchy. The most appropriate representation would be to conclude that Hippo signaling acts on both Bmp signaling and Hand2 expression. These could be independent influences, or they could be linked, but the data do not clearly support the direct linkage.

4) The authors should look through the manuscript carefully, as there are a number of errors in the text. Some examples, spotted by reviewers, are listed here:

- Instances where "specification" is used, when differentiation is appropriate

- "compared to humans" would read better as "compared to mouse and humans", given it follows a description of mice

- "transcriptional factor" should read "transcription factor"

- "restricts determines" is tautological and needs altering

- "frameshifts" should read "frameshift mutations"

- "faithfully recapitulates isl1 expression in vivo" is an overstatement if all isl1 expression was not assessed.

- “Interestingly, the gap length of WT embryos was significantly shorter than that of the lats1wt/ncv107lats2ncv108 embryos, the lats1/2 DKO embryos and the lats1/2 morphants.” However, the data in Figure 5 shows that gap length in wt embryos is larger.

5) It would be beneficial for the authors to review their choices of cited articles. In some cases, it seems as if the citations may not be the best choices to fully support the statements made. For example, does Hami et al. (2011) strongly support the points for which it is cited in the Introduction (that SHF cells come from the lateral and caudal ALPM, and that Isl1+ cells give rise to the inflow tract)?

https://doi.org/10.7554/eLife.29106.031

Author response

Essential revisions:

1) Importantly, the authors need to provide further evidence to support their conclusions regarding the differential fates of the left and right caudal ALPM. Previous cell tracking work has shown that cells from the left LPM form the dorsal side of the cardiac tube while cells from the right LPM form the ventral side of the cardiac tube (PMID: 18267096). Different from the description in the current manuscript, the older work showed that both the right and left LPM cells contributed to the venous pole. However, the dorsal side of the tube extends more anteriorly compared to the ventral side. Looking closely at the data provided by the authors (e.g. by playing the last 3 hours (equivalent of 2 secs) of their movie backward and forward rapidly), one gets the impression that the right caudal ALPM also contributes to the venous pole at the ventral side. Likely due to the tissue depth at the ventral surface, the fluorescence intensity of these cells is weaker and, therefore, difficult to track. To definitively state that the left and right sides contribute to the venous and arterial poles, respectively, the authors should provide higher resolution (and 3D) tracking of both the right and left ALPM cells. In addition, complementary lineage tracing experiments (using photoconversion or other cell labeling techniques) should be employed to follow the cells labeled in the right and left ALPM to their destination in the heart at 48 hpf.

As the reviewers critically commented, the contribution of both sides of the cells in the ALPM to venous pole should be carefully examined. We have performed additional cell tracking experiments that show that the cells in the caudal regions of both sides of ALPM move to the venous pole (New Figure 3A, 3B, and Video 1). The number of the cells from the left ALPM was greater than those from the right ALPM. The cells in the both sides of LPM move toward the cardiac disk. Those cells stay in the arterial pole. Therefore, the cells in the both sides of ALPM can become both venous and arterial pole cells. In the revised manuscript, we decline our previous conclusion that the cells in the left and right ALPM become venous and arterial pole cells, respectively. Instead, we state that both sides of ALPM cells become the cells in the venous pole. Initially, we tried to understand the origin of the IFT of the atrium. Therefore, we still focus on the venous pole cells that give rise to the IFT cells.

In addition, following the reviewer’s advice, we have performed the photoconversion experiments using the embryos transiently injected with hand2BAC:tdEosFP and photoconverted at 12 ss. We have confirmed that both the left and right ALPM cells are localized in the inflow tract (IFT) of the atrium at 52 hpf (New Figure 3D). Therefore, we have concluded that the cells in the caudal region of the left ALPM cells contribute to venous pole and inflow tract development more than those in the right ALPM. These results are consistent with the previous report, although it is worth mentioning that more cells in the left ALPM move to the venous pole than those in the right ALPM.

2) The authors highlight the expression of the Tead reporter in the left caudal ALPM, but it is not clear whether or not this reporter is expressed in the right caudal ALPM. This needs to be clarified.

We admit that we should clarify the expression of Tead reporter in the both sides of ALPM. To examine whether the expression of Tead reporter is observed in the right side of the caudal region of the ALPM, we crossed TgBAC(hand2:GFP) with Tg(eef1a1l1:galdb-hTEAD2ΔN-2A-mCherry);Tg(uas:mRFP1). We found that Tead reporter is expressed in both the left and right sides of entire ALPM (New Figure 3E). Collectively, our new data demonstrate that Hippo signaling functions in the entire ALPM. Therefore, this data suggests – together with the results showing the hand2 promoter-activated cells – that Hippo signaling in the ALPM contribute to the hand2-positive cell formation in the ALPM.

3) The ectopic hand2 expression and bmp2 expression in lats1/2 mutants is bilateral and not restricted to the left ALPM (Figures 5 and 6). How does this reconcile with the authors' suggested laterality of Hippo signaling (as shown in their model in Figure 7)?

As the reviewers were concerned, given that the results of new experiments suggest that Hippo signaling functions in bilaterally, we have declined our previous conclusion and propose that hippo signaling functions in both the left and the right ALPM (New revised Figure 7) in the revised manuscript. This conclusion is consistent with our previous data showing the bilateral expression of both hand2 and bmp2b in the ALPM as the reviewers pointed out in the lats1/2 mutants.

4) In Figure 6, the authors show enhanced expression of bmp2b in lats1/2 mutants and loss of bmp2 expression in yap/wwtr mutants. The effects are visible in the entire ALPM and head region, while their data shown in Figure 3 suggests that Yap1 is only active in the posterior ALPM. This discrepancy is not addressed. How do the authors fit these data with their model in which Hippo signaling activity is different in the posterior versus anterior ALPM?

In the previous experiments, we just focused on the caudal region of the ALPM using the certain layer cells as the reviewers pointed out. We admit that we should carefully track the entire ALPM cells and the expression of Tead reporter as in the reply to the comments (1) and (2). In new Figure 3E and new Figure 3—figure supplement 1A, Tead expression is found in the entire ALPM but not in the rostral PLPM. Therefore, these results are consistent with the results of the enhanced expression of bmp2b in lats1/2 mutant and loss of bmp2b expression in yap1/wwtr1 mutants at the same developmental stage (Figure 6B), suggesting that Hippo signaling restricts the expression of bmp2b in the ALPM. These results were included in the model of new Figure 7.

5) It's possible that an increase in signaling through either pathway (Hippo or Bmp) would result in increased cells at the IFT. Is there increased TEAD reporter activity following bmp2b heat-shock?

Exactly, the number of the cells at the venous pole correlates with that of the IFT. Indeed, the results shown in Figure 1 prompted us to examine the cell number at the venous pole that is determined by Hippo signaling. We have demonstrated that both Tead and Bmp reporters are activated at the venous pole at 24 hpf (Figure 1—figure supplement 2B, and Figure 6E).

According to the reviewers’ suggestion, we have over-expressed bmp2b by the injection of hsp70l:bmp2b-2A-mCherry in the Tead reporter embryos. In line with the previous results, bmp2b expression did not affect Tead reporter activity (New Figure 6—figure supplement 2A). This new data supports our model that Hippo signaling functions upstream of Bmp-Smad signaling.

6) To reduce BMP signaling, the authors inject mRNA for smad7 (inhibitory Smad). It is not clear why the authors try to block BMP signalling during early stages of development since BMP signaling is required for proper dorsal-ventral patterning. Did the authors observe DV patterning defects in these experiments?

We admit that we did not have any better method to spatio-temporally block BMP signaling in the ALPM. Therefore, smad7 mRNA injection was the only possible way for us to inhibit BMP signaling (Figure 6G and H). We have examined the phenotype of general inhibition of Bmp signaling, dorsalized development. As shown in the New Figure 6—figure supplement 3A and 3B, injection of smad7 mRNA dose-dependently increased dorsalization of the embryos. Therefore, we conclude that smad7 mRNA can block Bmp-Smad signaling to examine the Bmp-dependent signal for hand2 expression.

7) The authors imply that lats1 and lats2 have a special influence on the Isl1-positive atrial cells and do not affect the numbers of the Isl1-negative atrial cells. However, this is not clearly demonstrated by the data presented here: the total number of atrial cells is presented at one stage, and the number of Isl1-positive cells is presented at a substantially earlier stage. Does the change in the number of Isl1-positive cells account for the entirety of the difference in atrial size? Similarly, is the impact of overexpressing bmp2b and/or adding DMH1 specific to the Isl1-positive population, or do these manipulations affect the entire atrium?

Unfortunately we cannot prove the direct relevance of the number of isl1-posoitve cells in the earlier stage to that of atrial cardiomyocytes in the later stage, because we have neither Tg line nor imaging system that allow us to 3-dimesionally track the cell fate from 20 hpf to 3 dpf. To show the direct contribution of Isl1-positive SHF cells to the formation of atrial cardiomyocytes, we further need to cell track of isl1 promoter-activated cells. In our data, the actual increase of the atrial CM was about 20 cells in the lats1/2 mutants at 74 hpf (Figure 1B). This increase paralleled the increase in the isl1 promoter-activated cells at 26 hpf (Figure 4E). In addition, the number of isl1 promoter-activated cell at 26 hpf and 96 hpf was about 25 cells (Figure 4C and 4E), suggesting that isl1-positive cells does not increase during this period, although we cannot exclude the possibility that isl1 promoter-activated cells become negative, while isl1-negative cells could become positive. At least, these data imply that Hippo signaling determines the number of isl1-positive cells that give rise to atrial cardiomyocytes.

We thank the reviewers for suggesting an experiment that supports our conclusions. According to the reviewers’ advice, we examined the effect of inhibition of Bmp signaling on isl1-positive SHF cells and isl1-negative cardiomyocytes. Inhibition of Bmp signaling resulted in a decrease in the number of isl1-positive cells but not in that of myl7-positive cardiomyocytes (New Figure 6I and 6J). Therefore, we assume that the increased number of atrial cardiomyocytes in the lats1/2 mutants is ascribed to the increased number of isl1-positive cells

8) The role of Yap1/Wwtr1 in promoting hand2 expression needs further clarification. The authors point to the reduction of hand2 expression in yap1;wwtr1 mutants, but it is unclear how specifically this phenotype relates to the regulation of hand2. Is the entire ALPM reduced in yap1;wwtr1 mutants? In addition, the relationship between the cardia bifida phenotype and the reduced hand2 expression in the yap;wwtr1 mutant should be more clearly articulated. The yap;wwtr1 mutant appears smaller than the control embryo shown. Corresponding bright field images would be helpful here.

Following the reviewers’ suggestion, we looked at the entire ALPM by simultaneous monitoring bright field images and confirmed that hand2 expression is significantly reduced in the entire anterior region (New Figure 2—figure supplement 2C). In the paper by the group of Prof. Yelon (Development, 2000), hand2 mutants (han) show a reduction of hand2 expression and exhibit cardia bifida, suggesting that the reduced hand2-dependent signaling leads to cardia bifida. The yap1/wwtr1 mutants showed reduced hand2 expression and cardia bifida (Figure 5D, new Figure 2—figure supplement 2C and 2D). We further found that the expression of nkx2.5, one of ALPM-derived FHF marker, was not affected in yap1/wwtr1 mutants (New Figure 2—figure supplement 2D). This data indicates that Hippo signaling is not involved in the differentiation of FHF-derived cardiomyocyte but inhibits Yap1/Wwtr1-dependent hand2 expression in the ALPM.

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

Essential revisions:

1) From the authors' data, it is clear that there is a positive relationship between Bmp signaling and the formation of venous pole cells. It is also clear that Hippo signaling has a negative influence on bmp2b expression. However, it is not yet clear whether the caudal boundary of hand2 expression in the ALPM is set by Hippo signaling repressing bmp2b expression at that location. While it may be challenging to test a model that relates Hippo signaling to bmp2b expression directly, it would be beneficial for the authors to clarify aspects of their story that are not consistent with that model. For example, while bmp2b expression is increased in lats1;2 mutants, it does not appear to be expanded, and this seems to contradict the authors' model. Can the authors clarify this, or adjust their model accordingly?

We agree with the reviewer that this needs clarification. bmp2b expression is actually very difficult to quantify and we did not obtain conclusive results when measuring its expression domain. Due to the presence of the eye near the bmp2b expression domain and the necessity of dissecting the yolk, the measure of the expression domain highly variable from embryo to embryo. However, we found that bmp2b expression levels in the lats1/2 mutants examined by in situ hybridization at 10 somite-stage (ss) were greater than that of the control (Figure 6B). We further show that bmp2b expression was increased two-fold in lats1/2 morphants compared to controls via qPCR performed at 24 hpf (see Author response image 1).

Bmp2b being a secreted protein, we hypothesize that increased Bmp2b expression levels might lead to an expanded BMP signaling domain as a result of an increased protein accumulation, thus leading to the activation of hand2 expression more caudally in the ALPM. Since this hypothesis remains speculative at this point, we eliminated the previous statement “bmp2b expression is induced by Lats1/2-Yap1/Wwtr1 signaling in the rostral region of PLPM” in the revised manuscript.

Author response image 1
Quantitative-PCR analyses of expression of bmp2b mRNAs in the whole embryos at 24 hpf injected with the lats1/2 MOs (n=4).

Relative expression of mRNA in the morphants to that of the control is calculated.

https://doi.org/10.7554/eLife.29106.028

We changed the model accordingly by deleting the description of cell autonomous signaling because we cannot exclude the possibility that BMP secreted from the ALPM cells might affect BMP signaling in a non-cell autonomous manner.

2) The authors conclude that Yap1 activity in the ALPM induces Bmp signaling and that Bmp signaling activates hand2 and Isl1 expression, but this conclusion is too simple. Published data in zebrafish have shown that Bmp signaling is upstream of both hand2 and nkx2.5. (Both are lost in Bmp signaling mutants.) Yap1/wwtr1 double mutants still have hand2 expression (although reduced) and normal nkx2.5 expression. The authors do not address whether Bmp signaling is affected in the Yap1/wwtr1 mutants using the bre:gfp reporter line (they only show that bmp2b expression is reduced in the ALPM). Neither do they try to rescue the Yap1/wwtr1 phenotype with Bmp overexpression. The proposed model suggesting that Yap1 signaling is upstream of Bmp signaling is therefore an oversimplification and needs adjustment.

The reviewers are right; this point deserves clarification. A possible explanation for the discrepancy of Bmp2b-dependent transcription of both hand2 and nkx2.5 between the previous reports and our results is that Yap1/Wwtr1-Bmp2b downstream signal varies spatially and temporally to regulate heart formation. We believe that the downstream targets of Bmp-mediated signal, especially the transcription factors (Smads)-dependent signal, promote distinct targets depending on their time of action. In support of this possibility, the Tead reporter’s activation is observed in the whole ALPM at 10 ss (New Figure 4E), whereas it was only active in the cells of the venous pole at 24 hpf (Figure 1—figure supplement 2B).

In order to address this critical point further, we performed additional experiments to define more precisely whether Yap1/Wwtr1 signaling is upstream of Bmp signaling. We followed the editor’s suggestion and studied the Tg(BRE:GFP) fish in the yap1/wwtr1 heterozygous allele. Although we tried to incross the yap1/wwtr1 heterozygous fish with the Tg(BRE:GFP) reporter, we could not get any viable embryos because the eggs were not fertilized. After many attempts, we concluded that our females were not fertile, possibly because of the presence of the Tg(BRE:GFP) transgene. Therefore, we alternatively investigated the downstream signaling of Bmp by performing immunostainings for phosphorylated Smad1/5/9 in yap1/wwtr1 mutant, lats1/2 mutant, and wild-type embryos (New Figure 6—figure supplement 1A). In the yap1/wwtr1 double mutant embryos, phosphorylated Smad1/5/9 was decreased, while in the lats1/2 double mutant embryos phosphorylated Smad1/5/9 was increased. These new data suggest that Hippo signaling in the ALPM suppresses Bmp signaling and confirm our previous conclusions.

In addition, following the reviewers’ advice, we tried to investigate whether injection of bmp2b mRNA rescues the yap1/wwtr1 mutant phenotypes. As bmp2b overexpression leads to ventralization, which is not compatible with the analysis of the bmp2b expression domain, we established the optimal concentration of bmp2b to inject in the embryo leading to significantly low ventralized phenotype. We found that 30% of the embryos showed the ventralized phenotype when using 1.5 pg of bmp2b (as shown in Author response image 2). When injected in mutant embryos, we could not detect a clear rescue of the global yap1/wwtr1 mutant morphology. Since the number of double mutants is only 1 out of 16, the number of embryos we could analyze was too low to directly assess the ALPM phenotype. Therefore, we could not draw definitive conclusions based on these experiments. This experiment requires a more complex approach such as tissue-specific expression of bmp2b in the ALPM using gata4 or nkx2.5 promoter in the double mutant. Unfortunately, providing these experiments in the time frame of the revision agreed with the editor is not possible since it requires the generation of new transgenic lines that are not available at the moment.

In light of the fact that nkx2.5 expression is not altered in the yap1/wwtr1 (as well as in the lats1/2 morphants), we agree with the reviewers that our model needs adjustment. We have now changed the Discussion so that it is clear that it is possible that the expression of nkx2.5 might be regulated by the Bmp signaling that is not downstream of the Hippo signaling. The fact that dorso-ventral axis formation is not solely dependent on the Yap1/Wwtr1-Bmp signaling supports the possibility for additional regulators of BMP signaling in the ALPM. Overall, we think our proposition that the Hippo signaling-dependent Bmp signaling acts through Hand2 expression to restrict ALPM cell fate to become venous pole cells is valid. The identity of pathways involved and their timing of action is an active line of research in the lab and will require more time to reach conclusions.

Author response image 2
Bright field images of the embryo injected with bmp2b mRNA (1.5 pg) and yap1ncv117wwtr1ncv114 embryo at 22 hpf.

Lateral view, anterior to the left. bmp2b mRNA leads to ventralized phenotype (left panel). Yap1/wwtr1 mutant exhibits the defect of posterior body elongation (right panel).

https://doi.org/10.7554/eLife.29106.029

3) The phenotype shown for an embryo overexpressing smad7 mRNA does not seem to be a standard BMP-induced dorsalization phenotype, which is surprising. The appearance of the embryo in Figure 6–figure supplement 3A and the fragmented cells in Figure 6H suggest that the overexpression of smad7 may be causing cell death. Additionally, the cardiac phenotype attained after smad7 overexpression was not quantified, weakening the conclusions made from this experiment. If the authors intend to demonstrate cell-autonomous epistasis, they will likely need to use another approach. (Perhaps overexpression of a dominant-negative Bmp receptor?)

We sincerely apologize for the confusion. It is due to the fact that the phenotype we obtained was variable and chose to display the mild phenotype embryos to exemplify dorsalized phenotype in the embryos injected with smad7 mRNA in the previous figure. We replaced the previous figure with a new supplemental figure (Figure 6—figure supplement 3A) showing a dorsalized embryo following the smad7 mRNA Injection. We quantified again the percentage of dorsalized-phenotype and this time only embryos with phenotypes shown in Figure 6—figure supplement 3B were considered dorsalized embryos. The number of dorsalized embryos is decreased by comparison to the previous figure but this does not alter our conclusions that 100 pg is the best concentration to perform the overexpression experiment in the Tg(isl1:GFP);Tg(myl7:Nls-mCherry) embryos.

As suggested by the reviewers, we now provide a revised version where we quantified the number of isl1+ SHF cells in the embryo injected with smad7 mRNA (new Figure 6F). While the number of isl1+ SHF cells was decreased by the injection of smad7 mRNA, we could not detect any fragmented cells by confocal microscopy of the heart region highlighted by the TgBAC(isl1:GFP);Tg(myl7:Nls-mCherry) reporters at 26 hpf (see Author response image 3). These results indicate that the decrease of the isl1+ SHF cells of the embryo injected with smad7 mRNA was due to the inhibition of Bmp signaling and not cell fragmentation. These results are consistent with our previous results showing the decrease of the isl1+ SHF cells treated with the BMP inhibitor DMH1 (Figures 6I and 6J).

Following the reviewers’ comment, we toned down our previous statement about cell- autonomous epistasis because Bmp is a secretory molecule and we cannot exclude non-cell autonomous regulation.

4) In the experiment using DNA injection to induce mosaic Bmp2b expression (Figure 6—figure supplement 2A), the cells expressing bmp2b-2A-mCherry look fragmented, suggesting toxicity. The tissue on the left side also looks necrotic and the Yap1 reporter is very weak, while the authors state that it is unaffected. These results are not appropriate for inclusion in the manuscript. It is unclear why the authors did not use the available hsp70:bmp2b transgenic line to generate mosaic embryos through transplantation.

We agree with the reviewer that the cells look fragmented. We performed new experiments by imaging the bmp2b positive cells using confocal microscopy at high magnification and these experiments confirmed that a significant fraction of the cells expressing bmp2b-2A-mCherry is fragmented. We thank the reviewers for noticing this oversight. As the reviewers suggested, the data have been deleted from the revised version. We did not proceed further with this approach, we thus decided to delete the previous Figure 6C, 6D, and Figure 6—figure supplement 2A.

Author response image 3
The number of fragmented isl1 promoter-activated SHF cells of the embryos injected with the smad7 mRNA is very low as indicated at the bottom.

The heart region was highlighted by the TgBAC(isl1:GFP);Tg(myl7:Nls-mCherry) embryos at 26 hpf (n=3).

https://doi.org/10.7554/eLife.29106.030

Nevertheless, we strongly believe that our conclusions are still valid in absence of these experiments. Collectively, we now have the following data supporting our claim that Bmp2b is regulated by Hippo signaling to determine the number of SHF cells in the venous pole.

1) The Hippo signaling pathway inhibits phosphorylation of Smad1/5/9 (New Figure 6—figure supplement 1A).

2) The forced expression of smad7 mRNA results in a decrease of SHF cells (New Figures 6E and 6F).

3) DMH1 treatment leads to a decrease of SHF cells (Figures 6I and 6J).

4) Both the activity of the Bmp reporter and the phosphorylation of Smad1/5/9 are increased in the venous pole (New Figures 6C, 6D, and Figure 6—figure supplement 2B).

Therefore, we feel confident that the data obtained by transient and mosaic expression of Bmp2b using Tol2 system are unnecessary to support our claim.

5) In their revised manuscript, the authors conclude that venous pole cells originate on both sides of the ALPM, but they also state that "more cells moved from the left ALPM to the venous pole compared to the cells coming from the right ALPM". It is not clear that this semi-quantitative comparison is warranted, since it does not seem as if the authors tracked all of the cells on both sides of the ALPM. In Video 1, the authors compare five cells on the left to two cells on the right and state that this is representative of six independent experiments. How do the numbers compare among the six experiments? Is this a quantitatively reproducible observation? Do the photoconversion experiments (Figure 3D) reinforce the view that more venous pole cells come from the left than from the right? It is challenging to make a quantitative conclusion from this type of mosaic analysis. The authors should clarify the basis for this quantitative comparison between the left and right sides, or, alternatively, adjust their interpretation. Additionally, it would be helpful if the authors could clarify what criteria they used to define whether or not a tracked cell (or a photoconverted cell) became part of the venous pole in these experiments. Do the seven tracked cells in Video 1 account for the entirety of the venous pole?

We agree with the reviewers’ comment that it is challenging to make quantitative conclusions about the proportion of cells coming from the left and right ALPM. We also agree that we cannot conclude about a potential bias between left and right contribution of ALPM cells into the SHF. It was a mistake on our side and we sincerely apologize for it. In fact, re-examining our data we find that the Tead reporter is bilaterally expressed equally between the two embryonic sides, which would be difficult to reconcile with our previous conclusion. Similarly, bmp2b expression is symmetrically altered in the lats1/2 mutants and morphants. We thus feel that exploring the asymmetric contribution of ALPM cell to the SHF is not in the focus of our study and unnecessary. We, therefore, decided not to include this conclusion and changed the previous statement “the more cells in the left ALPM migrate toward the venous pole cells” into “hand2 positive cells of venous pole differentiate from the caudal ALPM”.

6) The authors have revised their title and Abstract to accompany the changes to their model that they have incorporated into their revised manuscript. It may be beneficial to consider revising these further in order to better emphasize the key points that are most strongly supported by the current text. For example, the title claims that Hippo signaling "suppresses the differentiation of cardiac precursor cells" but it is not clear that suppression of differentiation is shown here. More venous pole cells are produced when Hippo signaling is inhibited, but it is not evident whether this occurs through alteration of differentiation (as opposed to alteration of specification or proliferation). To make this claim, the authors would need to provide additional data in support of this conclusion. Also, the authors emphasize that the activity of Hippo signaling occurs "prior to establishment of the heart field", but it is not clear what they mean by this phrase. When do they consider the heart field to be established, and what is the significance of its establishment as it relates to the conclusions of their manuscript? Overall, it would be more effective if the title and Abstract emphasized formation of the venous pole specifically, as that seems to be the focus of the manuscript.

We agree that the previous title, Abstract and model did not fully reflect our results as the editor pointed out. We changed the title and revised the Abstract in accordance with our results, which demonstrate the involvement of Hippo signaling in the LPM and subsequent venous pole formation. We agree that the involvement of Hippo signaling specifically in the heart field is not established in our study. Therefore, we changed the previous title into a more precise title with a focus on the venous pole specification. The new title is “Hippo signaling determines the number of atrial cells that originate from the anterior lateral plate mesoderm in zebrafish”.

We also agree that the heart field was not well introduced leading to confusion about the origin of the additional cardiomyocytes observed in the Hippo pathway mutants. We rewrote extensively the Introduction and the Results to clarify this important issue.

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

The manuscript has been improved through its second round of revision, both by adding new data and by removing less convincing elements. Altogether, the revisions have streamlined and strengthened the overall clarity of the message regarding the impact of the Hippo signaling pathway on the number of venous pole cells in zebrafish, while toning down some of the more speculative or inconsistent aspects. However, there are some remaining issues that need to be addressed, as outlined below:

1) With each revision of this manuscript, the authors have modified their title and Abstract to accompany the changes in their message. Further modification would help the current title to fit more closely with the main point of the paper, which revolves around the pathways regulating the number of venous pole (Isl1+) cells. The new title and Abstract seem to emphasize the whole atrium, even though the take-home message of the data is focused on the venous pole. Further adjustment is needed to align the title and Abstract with the rest of the manuscript.

We thank the editors and reviewers for the helpful suggestions. We changed the title and Abstract to focus on the venous pole as they pointed out. The new title is “Hippo signaling determines the number of venous pole cells that originate from the anterior lateral plate mesoderm in zebrafish”. In the Abstract, we changed the final sentence to “Hippo signaling defines venous pole cardiomyocyte number by […]” to focus on the venous pole.

2) The data that are provided to argue that overexpression of smad7 (as in Figure 6) is non-toxic are not convincing. The authors have undertaken control experiments, examining whether embryos injected with smad7 mRNA look healthy, but this context is quite different from the mosaic scenario presented in the experiments, in which three different mRNAs are co-injected. It remains unexplored whether this cocktail is causing cell death (and therefore loss of hand2 expression) and therefore no conclusion can be drawn from this experiment. Therefore, the experiments utilizing smad7 overexpression should be removed from the manuscript.

Following the comments, to avoid the ambiguity of our results, we deleted the mosaic experiment results (Former Figure 6 G and H). Therefore, we renewed Figure 6.

3) It appears premature for the authors to conclude a direct, linear relationship connecting Hippo signaling, Bmp signaling, and Hand2 expression. Without a formal epistasis experiment (e.g. DMH1 treatment in lats1/2 LOF and measuring the hand2 response), it does not seem appropriate to suggest this hierarchy. The most appropriate representation would be to conclude that Hippo signaling acts on both Bmp signaling and Hand2 expression. These could be independent influences, or they could be linked, but the data do not clearly support the direct linkage.

We agree with the reviewer’s comment. We cannot exclude the possibility that Hippo signaling directly regulates hand2 expression and cannot conclude that hand2 is downstream of Bmp2 signaling. Therefore, we added a sentence to clarify this in the Discussion and edited Figure 7. In the new Figure 7, the two arrows denote the possibility that Hippo signaling acts on both Bmp signaling and Hand2.

4) The authors should look through the manuscript carefully, as there are a number of errors in the text. Some examples, spotted by reviewers, are listed here:

- Instances where "specification" is used, when differentiation is appropriate

- "compared to humans" would read better as "compared to mouse and humans", given it follows a description of mice

- "transcriptional factor" should read "transcription factor"

- "restricts determines" is tautological and needs altering

- "frameshifts" should read "frameshift mutations"

- "faithfully recapitulates isl1 expression in vivo" is an overstatement if all isl1 expression was not assessed.

- “Interestingly, the gap length of WT embryos was significantly shorter than that of the lats1wt/ncv107lats2ncv108 embryos, the lats1/2 DKO embryos and the lats1/2 morphants.” However, the data in Figure 5 shows that gap length in wt embryos is larger.

We thank the reviewers/editors for listing up our mistakes in the manuscript. We carefully checked the manuscript including the mistakes listed up and corrected other grammatical errors.

5) It would be beneficial for the authors to review their choices of cited articles. In some cases, it seems as if the citations may not be the best choices to fully support the statements made. For example, does Hami et al. (2011) strongly support the points for which it is cited in the Introduction (that SHF cells come from the lateral and caudal ALPM, and that Isl1+ cells give rise to the inflow tract)?

According to the reviewers/editors’ comments, we checked whether the citations match our statements. When we found the best manuscript that fit our description, we replaced previous citations with new citations.

https://doi.org/10.7554/eLife.29106.032

Article and author information

Author details

  1. Hajime Fukui

    1. Department of Cell Biology, National Cerebral and Cardiovascular Center Research Institute, Suita, Japan
    2. University of Strasbourg Institute for Advanced Study (USIAS), Strasbourg, France
    3. Institut de Génétique et de Biologie Moléculaire et Cellulaire, Illkirch, France
    4. Centre National de la Recherche Scientifique, Illkirch, France
    5. Institut National de la Santé et de la Recherche Médicale, Illkirch, France
    6. Université de Strasbourg, Illkirch, France
    Contribution
    Conceptualization, Resources, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Writing—original draft, Project administration
    Competing interests
    No competing interests declared
    ORCID icon 0000-0002-7652-2222
  2. Takahiro Miyazaki

    Department of Cell Biology, National Cerebral and Cardiovascular Center Research Institute, Suita, Japan
    Contribution
    Resources, Investigation
    Competing interests
    No competing interests declared
  3. Renee Wei-Yan Chow

    1. Institut de Génétique et de Biologie Moléculaire et Cellulaire, Illkirch, France
    2. Centre National de la Recherche Scientifique, Illkirch, France
    3. Université de Strasbourg, Illkirch, France
    4. Institut National de la Santé et de la Recherche Médicale, Illkirch, France
    Contribution
    Writing—review and editing
    Competing interests
    No competing interests declared
  4. Hiroyuki Ishikawa

    Department of Cell Biology, National Cerebral and Cardiovascular Center Research Institute, Suita, Japan
    Contribution
    Resources, Investigation
    Competing interests
    No competing interests declared
  5. Hiroyuki Nakajima

    Department of Cell Biology, National Cerebral and Cardiovascular Center Research Institute, Suita, Japan
    Contribution
    Resources, Validation, Investigation
    Competing interests
    No competing interests declared
  6. Julien Vermot

    1. Institut de Génétique et de Biologie Moléculaire et Cellulaire, Illkirch, France
    2. Centre National de la Recherche Scientifique, Illkirch, France
    3. Université de Strasbourg, Illkirch, France
    4. Institut National de la Santé et de la Recherche Médicale, Illkirch, France
    Contribution
    Supervision, Writing—review and editing
    Competing interests
    No competing interests declared
    ORCID icon 0000-0002-8924-732X
  7. Naoki Mochizuki

    1. Department of Cell Biology, National Cerebral and Cardiovascular Center Research Institute, Suita, Japan
    2. AMED-Core Research for Evolutional Science and Technology (AMED-CREST), Japan Agency for Medical Research and Development (AMED), Tokyo, Japan
    Contribution
    Conceptualization, Supervision, Funding acquisition, Project administration, Writing—review and editing
    For correspondence
    mochizuki@ncvc.go.jp
    Competing interests
    No competing interests declared
    ORCID icon 0000-0002-3938-9602

Funding

Ministry of Education, Culture, Sports, Science, and Technology (15H01221)

  • Hajime Fukui

Takeda Medical Research Foundation

  • Hajime Fukui
  • Naoki Mochizuki

Uehara Memorial Foundation

  • Hajime Fukui

Cell Science Research Foundation

  • Hajime Fukui

Japan Society for the Promotion of Science (16H02618)

  • Naoki Mochizuki

Japan Agency for Medical Research and Development (13414779)

  • Naoki Mochizuki

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

Acknowledgements

We thank DY Stainier for the TgBAC(hand2:GFP) fish; and M Sone, T Babazono, K Hiratomi, M Ueda, and S Toyoshima for their technical assistance.

Ethics

Animal experimentation: Animal experimentation: The experiments using zebrafish were approved by the institutional animal committee of National Cerebral and Cardiovascular Center (Permit number:17003) and performed according to the guidelines of the Institute.

Reviewing Editor

  1. Deborah Yelon, University of California, San Diego, United States

Publication history

  1. Received: May 30, 2017
  2. Accepted: May 26, 2018
  3. Accepted Manuscript published: May 29, 2018 (version 1)
  4. Version of Record published: June 11, 2018 (version 2)

Copyright

© 2018, Fukui et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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