1. Cell Biology
  2. Developmental Biology

Metabolic modulation regulates cardiac wall morphogenesis in zebrafish

  1. Ryuichi Fukuda  Is a corresponding author
  2. Alla Aharonov
  3. Yu Ting Ong
  4. Oliver A Stone
  5. Mohamed El-Brolosy
  6. Hans-Martin Maischein
  7. Michael Potente
  8. Eldad Tzahor
  9. Didier YR Stainier  Is a corresponding author
  1. Max Planck Institute for Heart and Lung Research, Germany
  2. Weizmann Institute of Science, Israel
https://doi.org/10.7554/eLife.50161
Share this article
Cite this article
  1. Ryuichi Fukuda
  2. Alla Aharonov
  3. Yu Ting Ong
  4. Oliver A Stone
  5. Mohamed El-Brolosy
  6. Hans-Martin Maischein
  7. Michael Potente
  8. Eldad Tzahor
  9. Didier YR Stainier
(2019)
Metabolic modulation regulates cardiac wall morphogenesis in zebrafish
eLife 8:e50161.
https://doi.org/10.7554/eLife.50161
  2. Download BibTeX
  3. Download .RIS

Abstract

During cardiac development, cardiomyocytes form complex inner wall structures called trabeculae. Despite significant investigation into this process, the potential role of metabolism has not been addressed. Using single cell resolution imaging in zebrafish, we find that cardiomyocytes seeding the trabecular layer actively change their shape while compact layer cardiomyocytes remain static. We show that Erbb2 signaling, which is required for trabeculation, activates glycolysis to support changes in cardiomyocyte shape and behavior. Pharmacological inhibition of glycolysis impairs cardiac trabeculation, and cardiomyocyte-specific loss- and gain-of-function manipulations of glycolysis decrease and increase trabeculation, respectively. In addition, loss of the glycolytic enzyme pyruvate kinase M2 impairs trabeculation. Experiments with rat neonatal cardiomyocytes in culture further support these observations. Our findings reveal new roles for glycolysis in regulating cardiomyocyte behavior during cardiac wall morphogenesis.

Introduction

During development, the heart undergoes a series of morphogenetic changes to form a functional cardiac wall structure (Moorman and Christoffels, 2003; Staudt and Stainier, 2012). The outer wall of the developing ventricle consists of compact layer cardiomyocytes (CMs), while the inner wall consists of complex muscular ridges, termed trabeculae, which facilitate efficient cardiac contraction and oxygenation of the cardiac wall prior to the formation of coronary vessels (Sedmera et al., 2000; Staudt and Stainier, 2012). Disruption of ventricular wall morphogenesis is associated with congenital cardiac malformations, the most common type of birth defects (Fahed et al., 2013), yet the cellular and molecular mechanisms regulating this complex process remain unclear.

Neuregulin (NRG)/Erb-b2 receptor tyrosine kinase (ERBB) 2/4 signaling has been shown to be essential for cardiac trabeculation. Nrg1, Erbb2 and Erbb4 knockout mice (Gassmann et al., 1995; Lee et al., 1995; Meyer and Birchmeier, 1995) and nrg2a (Rasouli and Stainier, 2017) and erbb2 (Liu et al., 2010) mutant fish fail to form trabeculae. ERBBs are members of the epidermal growth factor (EGF) receptor tyrosine kinase family. NRGs are expressed by the endocardium (Corfas et al., 1995; Meyer and Birchmeier, 1995; Grego-Bessa et al., 2007; Rasouli and Stainier, 2017) and bind to ERBBs on CMs, triggering homo- or heterodimerization of ERBB family members and leading to activation of downstream pathways (Sanchez-Soria and Camenisch, 2010). However, the targets of ERBB2 signaling that regulate CM behavior during trabeculation have not been identified.

Cardiac metabolism has been extensively studied in adult animals due to its central role in supplying energy for cardiac contraction (Doenst et al., 2013; Kolwicz et al., 2013). Adult CMs rely mostly on fatty acids as an energy substrate, and they are oxidized in mitochondria to generate ATP (Ellen Kreipke et al., 2016). Under conditions of hypertrophic or ischemic stress, CMs revert to glycolytic metabolism (Doenst et al., 2013), which is characteristic of embryonic cardiomyocytes and uses glucose as a fuel. Besides its role in energy generation, little is known about the role of metabolism during cardiac development.

Here, using high-resolution single cell imaging in zebrafish, we first show that developing CMs undergo extensive shape changes during the formation of the trabecular layer. By modulating glucose metabolism pharmacologically, we show that glycolysis regulates these processes. Using CM-specific loss- and gain-of-function models as well as mutant animals compromised in their glycolytic activity, we identify a role for glycolysis in cardiac wall morphogenesis. This study provides new insights into the role of cardiac metabolism in cardiac development.

Results

Cardiomyocytes that enter the trabecular layer exhibit distinct behaviors

During cardiac trabeculation in zebrafish and mouse, CMs delaminate from the compact layer to seed the trabecular layer (Liu et al., 2010; Zhang et al., 2013; Staudt et al., 2014; Jiménez-Amilburu et al., 2016; Del Monte-Nieto et al., 2018). Although CM behavior during trabeculation has been observed in zebrafish (Staudt et al., 2014; Cherian et al., 2016), the 3D morphology of single cardiomyocytes during the trabeculation process needs to be further explored. To this end, we performed 3D time-course imaging using chimeric hearts generated by cell transplantation. To label CM membranes and nuclei with EGFP and DsRed2 respectively, we used Tg(myl7:EGFP-HRAS); Tg(myl7:nDsRed2) cells as donors (Figure 1a and Figure 1—figure supplement 1a). We found that delaminating CMs exhibit morphological changes as well as rearrangements of contact sites (Figure 1b–c” and Figure 1—figure supplement 1b–d”; Figure 1—videos 1 and 2), while CMs remaining in the compact layer do not exhibit such changes (Figure 1d–e”). To examine cell-cell junctions during delamination, we analyzed N-cadherin (Cdh2), a major adherens junction component, at single cell resolution, and to this end used Tg(myl7:EGFP-HRAS); Tg(myl7:cdh2-tdTomato) cells as donors (Figure 1—figure supplement 1e). We observed that N-cadherin localizes to protruding membranes in delaminating CMs (Figure 1—figure supplement 1f–g”) and to the lateral membranes of compact layer CMs (Figure 1—figure supplement 1h–i”), in agreement with a previous report (Cherian et al., 2016). Next, we analyzed sarcomere structure during delamination using Tg(myl7:LIFEACT-GFP); Tg(myl7:nDsRed2) cells as donors. We found that CMs display partial sarcomere disassembly in their protrusions when entering the trabecular layer (Figure 1f–g”). These data indicate that delaminating CMs exhibit distinct behaviors including dynamic cell shape changes.

Figure 1 with 3 supplements see all
Download asset Open asset
Cardiomyocyte behavior during cardiac trabeculation.

(a) Schematic of the transplantation experiment. (b–e) 3D time-course images of chimeric hearts; magnified view (b’, c’, d’, e’) of area in white boxes and Y-Z plane images (b”, c”, d”, e”) along white dashed lines (b’, c’, d’, e’). CMs initially in the compact layer (b’, b”) enter the trabecular layer (c’, c”) exhibiting morphological changes and membrane protrusions (c’; arrowheads; n = 5 CMs); CMs remaining in the compact layer (d’, d”, e’, e”) do not exhibit obvious morphological changes (n = 5 CMs). The same CMs are shown at 126 and 175 hpf as indicated in the images. (f, g) 3D time-course images of chimeric heart; magnified view (f’, f”, g’, g”) of area in white boxes. CMs entering the trabecular layer exhibit partial disassembly of their sarcomeres (g’; arrowhead). Scale bars, 20 μm.

ERBB2 signaling activates glycolysis in cardiomyocytes

To gain additional insight into the molecular mechanisms that regulate trabeculation, we focused on ERBB2 signaling which is essential for this process in both mouse (Gassmann et al., 1995; Lee et al., 1995; Meyer and Birchmeier, 1995) and zebrafish (Liu et al., 2010; Peshkovsky et al., 2011; Rasouli and Stainier, 2017). In order to identify targets of ERBB2 signaling, we analyzed protein expression in a CM-specific transgenic mouse model inducibly expressing a constitutively active form of ERBB2 (CAERBB2) (D’Uva et al., 2015). We found that upon CAERBB2 overexpression (OE), several glycolytic enzymes were upregulated, while mitochondrial proteins and oxidative phosphorylation (OXPHOS)-related enzymes were downregulated (Figure 2—source data 1). We also tested the effects of Erbb2 OE, which like CAErbb2 OE, activates downstream signaling (Pedersen et al., 2009), on the expression of glycolytic enzyme genes in rat neonatal CMs, and found that many of them were upregulated (Figure 2a). Notably, we found that Erbb2 OE in rat neonatal CMs greatly upregulated the levels of pyruvate kinase M2 (PKM2), a key glycolytic enzyme, (Figure 2b and c), and increased glycolytic activity as evidenced by measuring extracellular acidification rate (ECAR) (Figure 2d). NRG1 stimulation also activated glycolysis in rat neonatal CMs (Figure 2—figure supplement 1a). These findings are also supported by a study analyzing changes in mRNA levels in caErbb2 OE mouse hearts (Honkoop et al., 2019). Moreover, we treated zebrafish embryos with an Erbb2 inhibitor (Figure 2e), which has been shown to severely affect trabeculation (Figure 2—figure supplement 1b) (Liu et al., 2010; Peshkovsky et al., 2011), and found that the cardiac expression of glycolytic enzyme genes was downregulated (Figure 2b). Altogether, these data indicate that ERBB2 signaling activates glycolysis in CMs.

Figure 2 with 2 supplements see all
Download asset Open asset
ERBB2 signaling activates glycolysis in cardiomyocytes.

(a) qPCR analysis of mRNA levels of glycolytic enzyme genes in control and Erbb2 overexpressing (OE) rat neonatal CMs (n = 3). Error bars, s.e.m. (b) Staining for PKM2, CTNI and DNA (DAPI) in control and Erbb2 OE rat neonatal CMs; arrowheads point to PKM2+ CMs. (c) Western blot analysis of PKM2 levels in control and Erbb2 OE rat neonatal CMs. (d) Extracellular acidification rate (ECAR) analysis in control and Erbb2 OE rat neonatal CMs; glycolytic capacity shown on the right (n = 7). Error bars, s.d. (e) qPCR analysis of mRNA levels of glycolytic enzyme genes in DMSO and Erbb2 inhibitor treated zebrafish hearts (n = 3). Error bars, s.e.m.; *p<0.05 and **p<0.001 by two-tailed unpaired t-test. NS, not significant. Scale bar, 20 μm.

Figure 2—source data 1

Mass spectrometry data.

P7 WT and CAERBB2 OE mouse hearts were isolated and protein expression levels analyzed by mass spectrometry. All presented proteins are statistically significant at p<0.05.

https://cdn.elifesciences.org/articles/50161/elife-50161-fig2-data1-v2.docx
Figure 2—source data 2

Primer sequences for qPCR analysis.

Primer sequences used in Figure 2a and e.

https://cdn.elifesciences.org/articles/50161/elife-50161-fig2-data2-v2.docx
Figure 2—source data 3

Mean Ct values of qPCR analysis in Figure 2a and e.

https://cdn.elifesciences.org/articles/50161/elife-50161-fig2-data3-v2.docx

Glycolysis regulates cardiomyocyte delamination during development

In order to analyze the role of glycolysis during trabeculation, we first focused on pyruvate metabolism. The pyruvate dehydrogenase complex (PDC) catalyzes the conversion of pyruvate to acetyl-coenzyme A (acetyl-CoA), which enters the tricarboxylic acid cycle (Zhang et al., 2014). Pyruvate dehydrogenase kinases (PDKs) inhibit PDC activity and enhance glycolysis in CMs (Zhao et al., 2008) and cancer cells (Koukourakis et al., 2005; Lu et al., 2008; Leclerc et al., 2017; Peng et al., 2018), thereby regulating the switch between glycolysis and OXPHOS (Zhang et al., 2014). Our analyses show that ERBB2 signaling positively regulates Pdk3 gene as well as protein expression (Figure 2a and b and Figure 2—source data 1). Thus, we hypothesized that PDK3 was one of the key enzymes regulating glycolysis in delaminating CMs in response to Erbb2 signaling. Consistent with this model, we found that the PDK inhibitor dichloroacetate (DCA) led to a significant reduction in the number of CMs in the trabecular layer (Figure 3a). Of note, this phenotype is similar to the one caused by Erbb2 inhibition (Figure 3a).

Figure 3 with 1 supplement see all
Download asset Open asset
Glycolysis regulates cardiac trabeculation.

(a) Confocal images (mid-sagittal sections) of 77 hpf hearts treated with DMSO, dichloroacetate (DCA) or Erbb2 inhibitor; magnified view of area in white boxes shown below; arrowheads point to CMs in the trabecular layer; percentage of CMs in the trabecular layer shown on the right (n = 5–7 ventricles). (b) Confocal images (mid-sagittal sections) of 77 hpf Tg(myl7:BFP-CAAX) alone or in combination with Tg(myl7:pdha1aSTA-P2A-tdTomato) or Tg(myl7:pdk3b-P2A-tdTomato) hearts; magnified view of area in white boxes shown below; arrowheads point to CMs in the trabecular layer; percentage of CMs in the trabecular layer shown on the right (n = 5–7 ventricles). (c–e”) Staining for CTNI, N-cadherin and DNA (DAPI) in control (c), Pdk3 (d) and Erbb2 (e) OE rat neonatal CMs; magnified view of area in yellow (c’, d’, e’) and white (c”, d”, e”) boxes; percentage of CMs exhibiting membrane protrusions shown on the right (n = 3 individual experiments; each value corresponds to an average of 30 CMs). Pdk3 and Erbb2 OE causes rat neonatal CMs to exhibit membrane protrusions (d’, e’; arrows) and cell-cell junction rearrangements (d’, e’; arrowheads). Error bars, s.e.m.; *p<0.05 and **p<0.001 by ANOVA followed by Tukey’s HSD test. Scale bars, 20 μm.

We next focused on pyruvate dehydrogenase E1 alpha 1 subunit a (pdha1a), which encodes a catalytic subunit of the PDC. Analysis of CM-specific loss of Pdha1 in mice has revealed the importance of PDC activity for OXPHOS (Sun et al., 2016). We generated a Tg(myl7:pdha1aSTA-P2A-tdTomato) line to overexpress an activated form of Pdha1a in CMs. This activated form of Pdha1a (Pdha1aSTA) contains mutations in its phosphorylation sites and thus is not inhibited by PDK. As a result, glycolysis is reduced and OXPHOS enhanced, as previously shown in cancer cells (Hitosugi et al., 2011; Fan et al., 2014). Notably, larvae expressing this activated form of Pdha1a exhibited a significant decrease in the number of CMs in the trabecular layer (Figure 3b and Figure 3—figure supplement 1a). We also generated a Tg(myl7:pdk3b-P2A-tdTomato) line to overexpress pdk3b in CMs and thereby promotes glycolysis (Lu et al., 2008), and found that these transgenic larvae exhibited a significant increase in the number of CMs in the trabecular layer (Figure 3b). Next, we tested whether the modulation of glycolysis affected CM proliferation and found that Tg(myl7:pdha1aSTA-P2A-tdTomato) or Tg(myl7:pdk3b-P2A-tdTomato) larvae did not exhibit a significant change compared to WT in the percentage of mVenus-gmnn+ CMs (Figure 3—figure supplement 1b). Furthermore, we examined whether the modulation of glycolysis affected CM morphology in rat neonatal CMs in culture and found that Pdk3 overexpression led to the induction of membrane protrusions (Figure 3d and d’), as well as cell-cell junction rearrangements (Figure 3d–d”). Similar effects were also observed following Erbb2 overexpression (Figure 3e–e”). Together, these data suggest that CM morphological changes regulated by glycolysis are important for delamination.

In order to further assess the role of glycolysis in trabeculation, we focused on zebrafish pkm2 to analyze a glycolytic enzyme mutant model. Loss of PKM2 has been shown to impair glycolysis in endothelial cells (Stone et al., 2018), and PKM2 expression has been associated with glycolysis and cell growth in cancer cells (Christofk et al., 2008). Moreover, in rat neonatal CMs, Erbb2 OE upregulated Pkm2 (Figure 2b and c). Mammalian Pkm encodes two splice variants (M1 and M2 isoforms); PKM2 plays an important role in glycolysis, while PKM1 promotes OXPHOS (Christofk et al., 2008; Lunt et al., 2015; Zheng et al., 2016). Zebrafish pkma2, a splice variant of pkma, and pkmb are the orthologues of mammalian Pkm2 (Stone et al., 2018). During early development, pkma is expressed in the heart, head, spinal cord and blood vessels, while pkmb is highly expressed in the somites (Figure 4—figure supplement 1a). At later stages, pkmb becomes clearly expressed in the heart (Figure 4—figure supplement 1b; Gunawan et al., 2019). We examined pkma2; pkmb double mutants in which Pkma1, which drives pyruvate metabolism via OXPHOS, remains intact, and found that loss of pkma2 and pkmb impaired trabeculation (Figure 4—figure supplement 1c). We did not find evidence for increased CM apoptosis in pkma2; pkmb double mutants compared to WT (Figure 4—figure supplement 1d). In order to examine the CM-specific role of pkma2 and pkmb in trabeculation, we performed cell transplantation experiments whereby pkma2; pkmb double heterozygous and double mutant cells were transplanted into WT embryos (Figure 4a). We found a significantly lower percentage of double mutant versus double heterozygous CMs in the trabecular layer of mosaic hearts (Figure 4b–c), indicating the importance of these genes in trabeculation. We also counted the number of trabecular CMs in these chimeric hearts and observed no significant deviation from WT (Figure 4—figure supplement 1e). Altogether, these results indicate that glycolysis plays important and CM-autonomous roles during trabeculation.

Figure 4 with 1 supplement see all
Download asset Open asset
Loss of pkm2 impairs cardiac trabeculation.

(a) Schematic of the transplantation experiment. (b, b’) 3D and mid-sagittal section images of chimeric hearts using pkma2+/-; pkmb+/-; Tg(myl7:EGFP-HRAS) (b) and pkma2-/-; pkmb-/-; Tg(myl7:EGFP-HRAS) (b’) cells as donors; magnified view of area in white boxes shown below. (c) Percentage of donor-derived trabecular CMs (n = 10 ventricles). Error bars, s.e.m.; *p<0.05 by two-tailed unpaired t-test. Scale bars, 20 μm.

Discussion

During trabeculation, CMs exhibit membrane protrusions (Staudt et al., 2014) and rearrange their cell-cell junctions (Cherian et al., 2016; Miao et al., 2019). Our 3D single CM imaging clearly reveals that CMs that enter the trabecular layer change their shape, similar to migrating cells, and lose cell-cell adhesion, indicating that they undergo phenotypic changes. Epithelial cells exhibit cellular plasticity as they change shape, and lose cell-cell adhesion and apicobasal polarity - a phenotypic transformation called epithelial to mesenchymal transition (EMT) (Nieto, 2013; Ye and Weinberg, 2015; Varga and Greten, 2017). Recent studies suggest that endothelial cells can also undergo phenotypic changes towards mesenchymal-like cells (Markwald et al., 1977; Zeisberg et al., 2007; Pearson, 2015; Dejana et al., 2017; Kovacic et al., 2019). Before the onset of trabeculation, compact layer CMs exhibit apicobasal polarity, and then some of them depolarize and subsequently delaminate to seed the trabecular layer (Jiménez-Amilburu et al., 2016). Notably, ERBB2 signaling, which is essential for trabeculation (Lee et al., 1995; Liu et al., 2010), induces EMT in breast cancer cells (Carpenter et al., 2015; Ingthorsson et al., 2016). Altogether, these data indicate that during trabeculation CMs undergo an EMT-like process triggered by ERBB2 signaling.

Cardiac metabolism is essential for energy production to sustain continuous cardiac contractions (Doenst et al., 2013). However, the role of metabolism during cardiac development remains unclear. Our study reveals that glycolysis regulates CM behavior during cardiac wall morphogenesis. Glycolysis enables rapid production of ATP to meet the high-energy demands of cell proliferation and migration in developing tissues, cancer cells and endothelial cells (Lunt and Vander Heiden, 2011; Liberti and Locasale, 2016; Potente and Carmeliet, 2017). Moreover, glycolytic intermediates are utilized to produce biomass including proteins and nucleic acids, further supporting these processes (DeBerardinis et al., 2008; Potente and Carmeliet, 2017). In addition, excessive glycolysis is associated with EMT in cancer cells (Peppicelli et al., 2014; Huang and Zong, 2017; Morandi et al., 2017). These findings indicate that glycolysis could regulate CM behavior in several different ways, and it will be interesting to dissect these processes further.

Materials and methods

Key resources table
Reagent type
(species) or resource		DesignationSource or referenceIdentifiersAdditional
information
Genetic reagent (Danio rerio)pkma2s717Stone et al., 2018
Genetic reagent (Danio rerio)pkmbs718Stone et al., 2018
Gene (Danio rerio)pdk3bNM_001080688
Gene (Danio rerio)pdha1aNM_213393
Gene (Rattus norvegicus)Pdk3NM_001106581
Gene (Homo sapiens)ERBB2NM_004448
Antibodyanti-Cardiac Troponin I (Goat polyclonal)AbcamAB_880622
Cat# ab56357		IF(1:500)
Antibodyanti-PKM2 (Rabbit monoclonal)Cell SignalingAB_1904096
Cat# D78A4		IF(1:100)
WB(1:1000)
Chemical compound, drugSodium dichloroacetateSigma Aldrich347795
Software, algorithmZen 2012 (Blue edition)Carl Zeiss MicroscopyVersion 1.1.2.0
Software, algorithmImaris x64BitplaneVersion 9.3.0
OtherDAPI stainSigmaD954(1 µg/mL)

Zebrafish

All zebrafish husbandry was performed under standard conditions in accordance with institutional (MPG) and national ethical and animal welfare guidelines. The following transgenic lines and mutants were used: Tg(myl7:EGFP-Has.HRAS)s883 (D'Amico et al., 2007) abbreviated Tg(myl7:EGFP-HRAS), Tg(myl7:LIFEACT-GFP)s974 (Reischauer et al., 2014), Tg(−5.1myl7:DsRed2-NLS)f2Tg (Mably et al., 2003) abbreviated Tg(myl7:nDsRed2), Tg(kdrl:Has.HRAS-mCherry)s896 (Chi et al., 2008) abbreviated Tg(kdrl:HRAS-mCherry), Tg(myl7:cdh2-tdTomato)bns78 (Fukuda et al., 2017), Tg(myl7:BFP-CAAX)bns193 (Guerra et al., 2018), Tg(myl7:mVenus-Gemnn)ncv43Tg (Jiménez-Amilburu et al., 2016), pkma2s717 (Stone et al., 2018) and pkmbs718 (Stone et al., 2018). To generate Tg(myl7:pdk3b-P2A-tdTomato)bns365 and Tg(myl7:pdha1aSTA-P2A-tdTomato)bns366, pdk3b (NM_001080688) and pdha1a (NM_213393) were isolated by RT-PCR and cloned under the control of the myl7 promoter in a vector containing Tol2 elements and two I-SceI restriction enzyme sites. The following primers were used to amplify the cDNA: pdk3b (forward 5’- AAGCAGACAGTGAACAAGCTTCCACCATGAAACTGTTTATCTGCCTACTG-3’ and reverse 5’-TAGCTCCGCTTCCGTCGACTCTGTTGACTTTGTATGTGGAC-3’); pdha1a (forward 5’-AAGCAGACAGTGAACAAGCTTCCACCATGAGAAAGATGCTAACCATAATT-3’ and reverse 5’-TAGCTCCGCTTCCGTCGACGCTGATGGACTTGAGTTTG-3’).

To generate the plasmid encoding an activated form of pdha1a (pdha1aSTA), the equivalent residues for human PDHA1 Ser293 and Tyr301 were replaced by alanine using the following primers: pdha1aSTA (forward 5’- CTATCGTTATCATGGACACGCTATGAGCGACCCAGGAGTCAGCGCCCGCACACGTGAGGAGA-3’ and reverse 5’- TTCCCTCACGTGTGCGGGCGCTGACTCCTGGGTCGCTCATAGCGTGTCCATGATAACGATAG-3’). Plasmids were then injected into one-cell stage embryos with I-SceI (NEB) or Tol2 mRNA.

Quantification of CMs in the trabecular layer

Request a detailed protocol

Quantification of trabecular CMs in 77 and 81 hpf heats was performed as previously described (Jiménez-Amilburu et al., 2016) using the ZEN software (ZEISS). Starting from the mid-sagittal plane, we quantified trabecular versus compact layer CMs in the ventricular outer curvature, 12 planes up and 12 planes down at an increment of 1 μm per plane. To quantify donor-derived trabecular CMs in chimeric hearts generated by cell transplantation, we counted the number of donor-derived trabecular and compact layer CMs. Then, the percentage of donor-derived trabecular CMs was calculated by dividing the number of donor-derived trabecular CMs by the total number of donor-derived CMs. Quantification of trabecular CMs in 131 hpf hearts was performed using the ZEN software (ZEISS). Starting from the mid-sagittal plane, we measured the whole myocardial area as well as the trabecular area, 10 μm up and 10 μm down. Three different sagittal planes per heart were measured. The percentage of the trabecular area for each sagittal plane was calculated by dividing the trabecular area by the myocardial area, and the average value was used for the graph.

In situ hybridization

Request a detailed protocol

In situ hybridization was performed as previously described (Thisse and Thisse, 2008). To synthesize pkma (NM_199333) and pkmb (NM_001003488) RNA probes, the following primers were used to amplify the corresponding DNA fragments: pkma (forward 5’- TTGGATCCACCATGTCTCAAACTAAAGCTC-3’ and reverse 5’- TTTGAATTCTTACGGCACTGGGACGACAC-3’); pkmb (forward 5’- TTGGATCCACCATGTCTCAGACAAAGACTA-3’ and reverse 5’- TTTGAATTCTCAAGGCACCACAACGATG’). The DNA fragments were cloned into the pCS2 vector. DIG-labeled RNA probes were synthesized using a DIG RNA labeling kit (Sigma-Aldrich) and MegaScript T7 Transcription Kit (Thermo Fisher Scientific).

Pharmacological treatments

Request a detailed protocol

Zebrafish embryos were treated with DMSO (control), 30 mM DCA (Sigma-Aldrich) or 5 μM Erbb2 inhibitor (AG1478; Sigma-Aldrich) from 50 hpf to 77 hpf and then analyzed.

TUNEL assay

Request a detailed protocol

To examine apoptosis, an in situ cell death detection kit (Roche) was used.

Immunostaining

Request a detailed protocol

Rat neonatal CMs were fixed in 4% paraformaldehyde. Anti-cardiac troponin I (CTNI) 1:500 (ab56357, Abcam) and anti-PKM2 1:100 (D78A4, Cell Signaling) were used. After washing with PBS, samples were stained with Alexa-568, Alexa-488 or Alexa-647 secondary antibodies 1:500 (Life Technologies), followed by 4′,6-Diamidine-2′-phenylindole dihydrochloride (DAPI) 1:2000 (Merck) staining to visualize DNA.

Cell culture

Request a detailed protocol

Rat neonatal (P2-P4) CMs were isolated and cultured as previously described (Fukuda et al., 2017). Cells were plated onto 0.1% gelatin-coated (Sigma) plates and cultured in DMEM/F12 (Gibco) supplemented with 5% horse serum, L-glutamine, Na-pyruvate, penicillin and streptomycin at 37°C and 5% CO2. Adenovirus vectors for transfection into CMs were generated using the AdEasy system (Agilent Technologies). To generate adenovirus vectors encoding PDK3 (NM_001106581) or ERBB2 (NM_004448), the following primers were used to amplify Pdk3 from rat neonatal CM or ERBB2 from Addgene clone # 39321: Pdk3 (forward 5’-TAGAGATCTGGTACCGTCGACCACCATGCGGCTCTTCTACCG-3’ and reverse 5’-GGATATCTTATCTAGAAGCTTCTAGAAAGTTTTATTACTCTTGATCTTGTCC-3’); ERBB2 (forward 5’-TAGAGATCTGGTACCGTCGACGCGGCCGCACCACCATGTATCCATATGATGTTCCAGATTATGCTATGGAGCTGGCGGCCTTG-3’ and reverse 5’-GGATATCTTATCTAGAAGCTTTCACACTGGCACGTCCAG).

Imaging

Request a detailed protocol

Zebrafish embryos and larvae were anesthetized with 0.2% tricaine and mounted in 1% low-melting agarose. A Zeiss spinning disk confocal microscope system (CSU-X1, Yokogawa) and ORCA-flash4.0 sCMOS camera (Hamamatsu) was used to acquire images. 3D images were processed using Imaris (Bitplane). Circularity was measured using ImageJ (NIH).

Western blotting

Request a detailed protocol

Protein expression levels were analyzed by western blotting as previously described (Fukuda et al., 2017). In brief, proteins were extracted with lysis buffer (150 mM Tris-HCl pH 7.5, 150 mM NaCl, 1% Triton X-100, 0.2% SDS, 1 mM EDTA, 5 mM NaF, 0.1 mM orthovanadate, 1 mM phenylmethylsulfonyl fluoride and 1 μg/ml aprotinin). Proteins were separated by SDS-PAGE. The following primary antibodies were used: anti-PKM2 1:1000 (D78A4, Cell Signaling) and anti-alpha-Tubulin 1:1000 (T6199, Sigma).

qPCR

Request a detailed protocol

Rat neonatal CMs were transfected with adenovirus vectors encoding genes of interest. 24 hr after transfection, total RNA was extracted. Zebrafish embryos and larvae were treated with DMSO or Erbb2 inhibitor from 55 to 106 hpf, and then the hearts were isolated to extract total RNA. A miRNeasy Mini kit (Qiagen) was used for total RNA extraction and cDNA was synthesized using a SuperScript Second Strand kit (Life Technologies). A CFX Connect Real-Time system (Bio-Rad) and DyNAmo colorFlash SYBR green qPCR kit (ThermoFisher Scientific) were used. Primer sequences are shown in Figure 2—source data 2 and Ct values in Figure 2—source data 3.

Metabolic assays

Request a detailed protocol

The extracellular acidification rate (ECAR) was measured using a Seahorse XFe96 analyzer (Seahorse Bioscience) following manufacturer’s protocol. Rat neonatal CMs were seeded at 30,000 cells/well on to 0.1% gelatin-coated XFe96 microplates (Agilent Technologies) in DMEM/F12 (Gibco) containing 10% fetal bovine serum (FBS), L-glutamine, Na-pyruvate, penicillin and streptomycin at 37°C and 5% CO2. After 24 hr of culture, the medium was replaced with serum-free medium. Then, cells were transfected with adenovirus vectors encoding genes of interest or a mock adenovirus vector, or treated with NRG1 (100 ng/ml; Abcam). 24 hr after transfection or NRG1 treatment, cells were maintained in non-buffered assay medium (Agilent Technologies) in a non-CO2 incubator 1 hr prior to the assay. A glycolysis stress test kit (Seahorse Bioscience) was used to monitor ECAR where baseline measurements were made followed by sequential injection of glucose (10 mM), oligomycin (2 μM), and 2-DG (100 mM).

Data availability

All data in this study are included in the manuscript and supporting files.

References

Decision letter

  1. Marianne E Bronner
    Senior and Reviewing Editor; California Institute of Technology, United States
  2. Shawn Burgess
    Reviewer; National Human Genome Research Institute, United States

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

This is an elegant study that asks multiple questions about the roles of metabolism in cardiac wall formation and the initiation of trabeculation. Cell transplantations between different genetic/transgenic backgrounds allows detailed analysis of the behavior of cells at the initiation of trabeculation. Overexpression of constitutively active ERRB2 (OE) resulted in upregulation of glycolytic pathway and activity in mouse transgenics, rat neonatal cardiomyocyte cultures, and downregulation of oxidative phosphorylation pathways; the converse was seen with inhibition of Errb2. Multiple approaches, including cardiomyocyte-specific expression of transgenics in zebrafish and pkma2; pkmb double mutants, indicate the importance of glycolysis for initiation of trabeculation, downstream of Errb2.

Decision letter after peer review:

Thank you for submitting your article "Metabolic modulation regulates cardiac wall morphogenesis and regeneration in zebrafish" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by Marianne Bronner as the Senior and Reviewing Editor. The following individual involved in review of your submission has agreed to reveal their identity: Shawn Burgess (Reviewer #1).

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

Although all of the reviewers are enthusiastic about the manuscript, they concur that it would be important to drop the regeneration component of the manuscript, which feels a bit added on and is not entirely convincing. Other changes are recommended but non-essential. The full reviews are included below for your information.

Reviewer #1:

Fukuda and colleagues have submitted a manuscript describing the role of metabolic shifts in cardiac development and regeneration. The majority of the work focuses on a particular developmental stage where the inner walls of the heart begin forming from the compact layer via delaminating cardiomyocytes (trabeculation). ErbB2 signaling has been shown to be important for this process and modulation of erbb2 activity levels is the starting point for this manuscript. Across three model systems, increases in ErbB2 activity increases genes related to glycolysis and decreases in ErbB2 signaling have matching decreases in genes involved in glycolysis. The authors targeted specific enzymes modulating pyruvate metabolism, a summary of the results are 1) inhibiting pyruvate dehydrogenase kinase decreases CMs, 2) increasing PDK activity increases CMs, 3) increasing pyruvate dehydrogenase activity decreases CMs, 4) decreasing pyruvate kinase decreases CMs. Shifting attention to cardiac regeneration in adult zebrafish, similar roles for metabolic shifts are seen, inhibition of PDK (with DCA) or phosphoglucose isomerase (2DG) reduces regeneration as does loss of PK activity while loss of ppargc1a (involved in mitochondrial biosynthesis) stimulates regeneration. In general, these are well-controlled experiments that add significantly to our understanding of the relationship between metabolic modulation and embryonic morphogenesis.

There are some problems with conclusions and/or conflicting data that should be addressed.

1) The authors have focused primarily on enzymes involved in the last steps of glycolysis: the pyruvate dehydrogenase complex (PDC), PDK, and pyruvate kinase (PK). It is true that these enzymes are listed in the glycolysis pathway, but except for PK which does release one ATP molecule, the other enzymes are primarily involved in shunting pyruvate either towards lactic acid (and the pentose phosphate pathway) or into the TCA cycle. This shift point isn't really modulating glycolysis but the choice between aerobic and anaerobic respiration and it may be that pyruvate concentration and/or utilization is the key step they are looking at and not glycolysis per se. I am arguing this point because otherwise the loss of PK activity (pkma2/pkmb) resulting in less trabeculation and regeneration appears to conflict with the other observations. Increasing PDK activity pushes pyruvate towards lactate (shunting away from mitochondria) resulting in an increase in trabeculation, blocking PDK activity has the opposite effect, less trabeculation more TCA cycle. The dominant active pyruvate dehydrogenase experiments are also consistent with these results, more PDK activity = less trabeculation. The PK knockouts should also reduce the availability of pyruvate for the mitochondria, yet it results in less trabeculation and less regeneration. Perhaps lactate is a key signaling molecule? This has been shown in neurons for example.

2) I think the authors need to change the general use of glycolysis throughout the manuscript to specify whether they are talking about anaerobic or aerobic glycolysis. Can the redox state be determined during trabeculation?

Reviewer #2:

There are a few questions about the trabeculation results, but overall this part of the study is convincing and an important contribution to the heart development field. In contrast, there are significant concerns about the heart regeneration aspects of the paper.

1) The interpretation that cardiac regeneration is altered is not convincing. The assays for regeneration were only taken to 5 days post cryo injury (dpci), which is not sufficient time to assess whether regeneration is normal or abnormal. The hearts in any of the experimental cohorts could be slightly delayed and then fully recovered by 30 dpci. There is an additional concern that the recovering fish are heat shocked every day for the first five days up to analysis. The standard in the field is 30- or 60-days post injury, with intermittent time points. I suggest removing the regeneration aspect of this paper. It is just too premature, unsupported conclusions in the title and Abstract could be incorrect.

2) In Figure 4, cell transplantation is used to generate chimeric hearts is a very useful experiment. I imagine the numbers are in hand, but it would be useful to see them analyzed in a few different ways. Are the total number of trabecular cardiomyocytes decreased in the hearts that received mutant donor cells? Or is it just a shift between the mutant vs. host cells that can contribute to the trabeculae? Figure 4C indicates that the% of cardiomyocytes that are trabecular was decreased in hearts that received homozygous mutant cells. However, it does not tell us whether the cardiomyocytes that are trabecular are from the transplanted or host cells. The legend says 'percentage of donor CMs' However, Figure 4B' appears to show some labeled (transplant) cells that are interior (trabecular), outside the white box. It would be useful to know the number of labeled (transplant) vs. unlabeled (host) cells that are trabecular, to understand whether the effect is cell autonomous or non-autonomous.

3) In Supplementary Figure 5, it appears that mCherry is also expressed in the epicardium after TAM and HS treatment. If this is the case, interpretation of results will need modification.

Reviewer #3:

In this manuscript, Fukuda et al. describe a role for glucose metabolism in not only normal development of the trabeculated myocardium, but also in the successful regeneration of the adult heart. Much is known about metabolic processes in the adult heart; here, the authors present a previously understudied role of glycolysis during development. First, by chimeric analysis to produce a higher-resolution look by live imaging, the authors show that cardiomyocytes elicit changes in cell shape to properly develop a trabeculated myocardium. Furthermore, the authors show Nrg1/Erbb2 signaling is required for trabeculation and activates glycolysis, which regulates development of the trabecular myocardium. Finally, the authors extend their study to adulthood and show that glycolysis is essential for successful dedifferentiation and proliferation of cardiomyocytes in cryoinjured adult hearts by upregulation of glycolytic enzymes at the wound border after injury. Overall, the authors contribute a new understanding of the role of glycolysis in proper heart development and repair.

1) The authors describe cell shape changes and membrane protrusions of cardiomyocytes to enter the trabecular layer. While the images are that of live embryos, this argument would be strengthened by time-lapse imaging to track the individual cardiomyocytes changing shape and position over time.

2) Methods used to determine glycolytic activity by assaying extracellular acidification rate should be explained in more detail.

3) The authors use transplanted double heterozygous and double mutant cells to show double mutant cells cannot incorporate into trabeculae. While these are very careful experiments, the health of these cells after transplantation is a concern. Have the authors scored transplant survival? Or performed a TUNEL assay to ensure the mutant cells are not dying after transplantation? Is delamination the only cellular event (cell motility, shape change, membrane protrusions, etc) that is perturbed?

4) Ppargc1a regulates genes involved in OXPHOS and are absent from regenerating hearts, but are they present in uninjured hearts by in situ?

https://doi.org/10.7554/eLife.50161.sa1

Author response

Reviewer #1:

[…] There are some problems with conclusions and/or conflicting data that should be addressed.

1) The authors have focused primarily on enzymes involved in the last steps of glycolysis: the pyruvate dehydrogenase complex (PDC), PDK, and pyruvate kinase (PK). It is true that these enzymes are listed in the glycolysis pathway, but except for PK which does release one ATP molecule, the other enzymes are primarily involved in shunting pyruvate either towards lactic acid (and the pentose phosphate pathway) or into the TCA cycle. This shift point isn't really modulating glycolysis but the choice between aerobic and anaerobic respiration and it may be that pyruvate concentration and/or utilization is the key step they are looking at and not glycolysis per se. I am arguing this point because otherwise the loss of PK activity (pkma2/pkmb) resulting in less trabeculation and regeneration appears to conflict with the other observations. Increasing PDK activity pushes pyruvate towards lactate (shunting away from mitochondria) resulting in an increase in trabeculation, blocking PDK activity has the opposite effect, less trabeculation more TCA cycle. The dominant active pyruvate dehydrogenase experiments are also consistent with these results, more PDK activity = less trabeculation. The PK knockouts should also reduce the availability of pyruvate for the mitochondria, yet it results in less trabeculation and less regeneration. Perhaps lactate is a key signaling molecule? This has been shown in neurons for example.

The mammalian Pkm gene gives rise to two splice isoforms, Pkm1 and Pkm2. The zebrafish genome contains two pkm genes, pkma and pkmb. pkma encodes both an orthologue of mammalian Pkm1 (pkma1) and an orthologue of mammalian Pkm2 (pkma2), while pkmb encodes only another orthologue of mammalian Pkm2 (Stone et al., 2018).

Expression of PKM2 is associated with metabolism of pyruvate to lactate. Loss of PKM2 has been shown to result in compensatory upregulation of PKM1, which drives enhanced mitochondrial metabolism of pyruvate (Christofk et al., 2008; Lunt et al., 2015; Zheng et al., 2016). Therefore, with respect to pyruvate metabolism, the loss of PKM2 mimics blocking PDK activity, whereas the loss of PKM1 mimics enhancing PDK activity. In pkma2; pkmb double mutant zebrafish, the pkma1 isoform, which drives pyruvate metabolism via OXPHOS, remains intact. Thus, in pkma2; pkmb double mutant zebrafish we expect a reduction in the metabolism of pyruvate to lactate and an increase in pyruvate oxidation, similar to the situation observed following inhibition of PDK activity. These findings are consistent with the other models that have been analyzed (Christofk et al., 2008; Zheng et al., 2016).

The revised sentence reads as follows:

‘We examined pkma2; pkmb double mutants in which Pkma1, which drives pyruvate metabolism via OXPHOS, remains intact, and found that loss of pkma2 and pkmb impaired trabeculation (Figure 4—figure supplement 1C).’

2) I think the authors need to change the general use of glycolysis throughout the manuscript to specify whether they are talking about anaerobic or aerobic glycolysis.

Although PKM2 (Christofk et al., 2008) and PDK (Takubo et al., 2013) play an important role during aerobic glycolysis (Vander Heiden et al., 2009; Lunt and Vander Heiden, 2011), we did not assess oxygen levels in our models, and thus prefer to use “glycolysis”.

Can the redox state be determined during trabeculation?

We have now examined the redox state using the cardiomyocyte specific reporter line previously used to examine hydrogen peroxide levels (Panieri et al., 2017). We found no significant difference in reporter activity between compact and trabecular cardiomyocytes during delamination (Author response image 1).

Author response image 1
Download asset Open asset
Redox analysis.

70 hpf Tg(myl7:mitochondrial-Rogofp2Orp1) hearts were examined and 405 nm/488 nm ratios determined for redox state analysis. Relative values of 405 nm/488 nm in compact and trabecular cardiomyocytes are shown.

Reviewer #2:

There are a few questions about the trabeculation results, but overall this part of the study is convincing and an important contribution to the heart development field. In contrast, there are significant concerns about the heart regeneration aspects of the paper.

1) The interpretation that cardiac regeneration is altered is not convincing. The assays for regeneration were only taken to 5 days post cryo injury (dpci), which is not sufficient time to assess whether regeneration is normal or abnormal. The hearts in any of the experimental cohorts could be slightly delayed and then fully recovered by 30 dpci. There is an additional concern that the recovering fish are heat shocked every day for the first five days up to analysis. The standard in the field is 30- or 60-days post injury, with intermittent time points. I suggest removing the regeneration aspect of this paper. It is just too premature, unsupported conclusions in the title and Abstract could be incorrect.

We have now removed the regeneration part from the revised manuscript.

2) In Figure 4, cell transplantation is used to generate chimeric hearts is a very useful experiment. I imagine the numbers are in hand, but it would be useful to see them analyzed in a few different ways. Are the total number of trabecular cardiomyocytes decreased in the hearts that received mutant donor cells? Or is it just a shift between the mutant vs. host cells that can contribute to the trabeculae? Figure 4C indicates that the% of cardiomyocytes that are trabecular was decreased in hearts that received homozygous mutant cells. However, it does not tell us whether the cardiomyocytes that are trabecular are from the transplanted or host cells. The legend says 'percentage of donor CMs.' However, Figure 4B' appears to show some labeled (transplant) cells that are interior (trabecular), outside the white box. It would be useful to know the number of labeled (transplant) vs. unlabeled (host) cells that are trabecular, to understand whether the effect is cell autonomous or non-autonomous.

We have now counted the total number of trabecular cardiomyocytes in the chimeric hearts and no significant differences were observed compared to WT (Figure 4—figure supplement 1D).

In Figure 4C, we counted the number of donor-derived cardiomyocytes in both the trabecular and compact layers of the chimeric hearts. We then calculated the percentage of donor-derived trabecular CMs by dividing the number of donor-derived trabecular CMs by the total number of donor-derived CMs. To clarify this point we have now modified Figure 4C, its legend and the relevant part of the Materials and methods section in the revised manuscript.

Together, our data indicate cardiomyocyte-autonomous function of pkma2 and pkmb during trabeculation.

3) In Supplementary Figure 5, it appears that mCherry is also expressed in the epicardium after TAM and HS treatment. If this is the case, interpretation of results will need modification.

We have now removed the regeneration part from the revised manuscript.

Reviewer #3:

[…] 1) The authors describe cell shape changes and membrane protrusions of cardiomyocytes to enter the trabecular layer. While the images are that of live embryos, this argument would be strengthened by time-lapse imaging to track the individual cardiomyocytes changing shape and position over time.

Time-lapse imaging to track individual cardiomyocytes during trabeculation has been shown in a previous paper (Staudt et al., 2014). However, the 3D morphology of single cardiomyocytes that enter the trabecular layer remains mostly unclear, which is why we examined it.

2) Methods used to determine glycolytic activity by assaying extracellular acidification rate should be explained in more detail.

We have now modified the Materials and methods section in the revised manuscript to address the reviewer’s comment.

3) The authors use transplanted double heterozygous and double mutant cells to show double mutant cells cannot incorporate into trabeculae. While these are very careful experiments, the health of these cells after transplantation is a concern. Have the authors scored transplant survival? Or performed a TUNEL assay to ensure the mutant cells are not dying after transplantation?

We have now performed TUNEL assays and no elevated levels of apoptosis were detected in pkma2; pkmb mutant cardiomyocytes in chimeric hearts compared to WT cardiomyocytes (Figure 4—figure supplement 1D).

Is delamination the only cellular event (cell motility, shape change, membrane protrusions, etc) that is perturbed?

NRG/ERBB signaling also regulates cardiomyocyte proliferation (Zhao et al., 1998; Grego-bessa et al., 2007; Bersell et al., 2009; Rasouli and Stainier, 2017). We have now counted the number of proliferating cardiomyocytes during trabeculation and these data indicate that modulation of metabolism does not appear to affect cardiomyocyte proliferation (Figure 3—figure supplement 1B).

4) Ppargc1a regulates genes involved in OXPHOS and are absent from regenerating hearts, but are they present in uninjured hearts by in situ?

We have now removed the regeneration part from the revised manuscript.

References

Bersell, K., Arab, S., Haring, B., and Kühn, B. (2009). Neuregulin1/ErbB4 Signaling Induces Cardiomyocyte Proliferation and Repair of Heart Injury. Cell 138, 257–270.

Panieri, E., Millia, C., Santoro, M.M., (2017). Real-time quantification of subcellular H2O2 and glutathione redox potential in living cardiovascular tissues, Free Radic Bio Med, 109, 189-200.

Takubo, K., Nagamatsu, G., Kobayashi, C.I., Nakamura-Ishizu, A., Kobayashi, H., Ikeda, E., Goda, N., Rahimi, Y., Johnson, R.S., Soga, T., et al. (2013). Regulation of glycolysis by Pdk functions as a metabolic checkpoint for cell cycle quiescence in hematopoietic stem cells. Cell Stem Cell 12, 49–61.

Vander Heiden MG, Cantley LC, Thompson CB. (2009). Understanding the Warburg effect: the metabolic requirements of cell proliferation. Science 324:1029–33.

Zhao, Y.Y., Sawyer, D.R., Baliga, R.R., Opel, D.J., Han, X., Marchionni, M.A., and Kelly,

R.A. (1998). Neuregulins promote survival and growth of cardiac myocytes: Persistence of ErbB2 and ErbB4 expression in neonatal and adult ventricular myocytes. J. Biol. Chem. 273, 10261–10269.

https://doi.org/10.7554/eLife.50161.sa2

Article and author information

Author details

  1. Ryuichi Fukuda

    Department of Developmental Genetics, Max Planck Institute for Heart and Lung Research, Ludwigstrasse, Germany
    Contribution
    Conceptualization, Project administration
    For correspondence
    Ryuichi.Fukuda@mpi-bn.mpg.de
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-0281-5161

  2. Alla Aharonov

    Department of Molecular Cell Biology, Weizmann Institute of Science, Rehovot, Israel
    Contribution
    Resources, Investigation
    Competing interests
    No competing interests declared

  3. Yu Ting Ong

    Angiogenesis & Metabolism Laboratory, Max Planck Institute for Heart and Lung Research, Ludwigstrasse, Germany
    Contribution
    Investigation
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-3407-2515

  4. Oliver A Stone

    Department of Developmental Genetics, Max Planck Institute for Heart and Lung Research, Ludwigstrasse, Germany
    Present address
    Department of Physiology, Anatomy and Genetics, BHF Centre of Research Excellence, University of Oxford, Oxford, United Kingdom
    Contribution
    Resources, Investigation
    Competing interests
    No competing interests declared

  5. Mohamed El-Brolosy

    Department of Developmental Genetics, Max Planck Institute for Heart and Lung Research, Ludwigstrasse, Germany
    Contribution
    Resources, Investigation
    Competing interests
    No competing interests declared

  6. Hans-Martin Maischein

    Department of Developmental Genetics, Max Planck Institute for Heart and Lung Research, Ludwigstrasse, Germany
    Contribution
    Investigation
    Competing interests
    No competing interests declared

  7. Michael Potente

    Angiogenesis & Metabolism Laboratory, Max Planck Institute for Heart and Lung Research, Ludwigstrasse, Germany
    Contribution
    Resources, Investigation
    Competing interests
    No competing interests declared

  8. Eldad Tzahor

    Department of Molecular Cell Biology, Weizmann Institute of Science, Rehovot, Israel
    Contribution
    Resources
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-5212-9426

  9. Didier YR Stainier

    Department of Developmental Genetics, Max Planck Institute for Heart and Lung Research, Ludwigstrasse, Germany
    Contribution
    Supervision
    For correspondence
    Didier.Stainier@mpi-bn.mpg.de
    Competing interests
    Senior editor, eLife
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-0382-0026

Funding

Max-Planck-Gesellschaft (Open-access funding)

  • Didier YR Stainier

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

Acknowledgements

We thank Beate Grohmann, Radhan Ramadass, Srinath Ramkumar, Simon Perathoner, Carmen Büttner, Nana Fukuda and Sharon Meaney-Gardian for help and support, Arica Beisaw, Ruben Marin-Juez, Josephine Gollin and Rashmi Priya for comments on the manuscript, and Jeroen Bakkers for communication. This work was supported in part by funds from the Max Planck Society to DYRS.

Ethics

Animal experimentation: All zebrafish husbandry was performed under standard conditions in accordance with institutional (MPG) and national ethical and animal welfare guidelines approved by the ethics committee for animal experiments at the Regional Board of Darmstadt, Germany (permit numbers B2/1017, B2/1041 and B2/1159).

Senior and Reviewing Editor

  1. Marianne E Bronner, California Institute of Technology, United States

Reviewer

  1. Shawn Burgess, National Human Genome Research Institute, United States

Publication history

  1. Received: July 12, 2019
  2. Accepted: December 20, 2019
  3. Accepted Manuscript published: December 23, 2019 (version 1)
  4. Version of Record published: February 4, 2020 (version 2)

Copyright

© 2019, Fukuda 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.

Metrics

  • 2,999
    Page views
  • 531
    Downloads
  • 15
    Citations

Article citation count generated by polling the highest count across the following sources: Crossref, PubMed Central, Scopus.

Download links

A two-part list of links to download the article, or parts of the article, in various formats.

Downloads (link to download the article as PDF)

Open citations (links to open the citations from this article in various online reference manager services)

Cite this article (links to download the citations from this article in formats compatible with various reference manager tools)

  1. Ryuichi Fukuda
  2. Alla Aharonov
  3. Yu Ting Ong
  4. Oliver A Stone
  5. Mohamed El-Brolosy
  6. Hans-Martin Maischein
  7. Michael Potente
  8. Eldad Tzahor
  9. Didier YR Stainier
(2019)
Metabolic modulation regulates cardiac wall morphogenesis in zebrafish
eLife 8:e50161.
https://doi.org/10.7554/eLife.50161

Categories and tags

Research organism

Further reading

    1. Developmental Biology
    2. Stem Cells and Regenerative Medicine

    Single-cell analysis uncovers that metabolic reprogramming by ErbB2 signaling is essential for cardiomyocyte proliferation in the regenerating heart

    Hessel Honkoop, Dennis EM de Bakker ... Jeroen Bakkers
    Research Article Updated

    While the heart regenerates poorly in mammals, efficient heart regeneration occurs in zebrafish. Studies in zebrafish have resulted in a model in which preexisting cardiomyocytes dedifferentiate and reinitiate proliferation to replace the lost myocardium. To identify which processes occur in proliferating cardiomyocytes we have used a single-cell RNA-sequencing approach. We uncovered that proliferating border zone cardiomyocytes have very distinct transcriptomes compared to the nonproliferating remote cardiomyocytes and that they resemble embryonic cardiomyocytes. Moreover, these cells have reduced expression of mitochondrial genes and reduced mitochondrial activity, while glycolysis gene expression and glucose uptake are increased, indicative for metabolic reprogramming. Furthermore, we find that the metabolic reprogramming of border zone cardiomyocytes is induced by Nrg1/ErbB2 signaling and is important for their proliferation. This mechanism is conserved in murine hearts in which cardiomyocyte proliferation is induced by activating ErbB2 signaling. Together these results demonstrate that glycolysis regulates cardiomyocyte proliferation during heart regeneration.

    1. Developmental Biology

    Heart Development and Regeneration: Metabolism makes and mends the heart

    Megan L Martik
    Insight

    Experiments in zebrafish have shed new light on the relationship between development and regeneration in the heart.

    1. Cell Biology

    Annexin A6 mediates calcium-dependent exosome secretion during plasma membrane repair

    Justin Krish Williams, Jordan Matthew Ngo ... Randy Schekman
    Research Article Updated

    Exosomes are an extracellular vesicle (EV) subtype that is secreted upon the fusion of multivesicular bodies (MVBs) with the plasma membrane. Exosomes may participate in intercellular communication and have utility as disease biomarkers; however, little is known regarding the physiological stimuli that induce their secretion. Ca2+ influx promotes exosome secretion, raising the possibility that exosomes are secreted during the Ca2+-dependent plasma membrane repair of tissues damaged by mechanical stress in vivo. To determine whether exosomes are secreted upon plasma membrane damage, we developed sensitive assays to measure exosome secretion in intact and permeabilized cells. Our results suggest that exosome secretion is coupled to Ca2+-dependent plasma membrane repair. We find that annexin A6 (ANXA6), a well-known plasma membrane repair protein, is recruited to MVBs in the presence of Ca2+ and required for Ca2+-dependent exosome secretion, both in intact and in permeabilized cells. ANXA6 depletion stalls MVBs at the cell periphery, and ANXA6 truncations localize to different membranes, suggesting that ANXA6 may serve to tether MVBs to the plasma membrane. We find that cells secrete exosomes and other EVs upon plasma membrane damage and propose that repair-induced secretion may contribute to the pool of EVs present within biological fluids.