1. Developmental Biology

Apelin signaling drives vascular endothelial cells toward a pro-angiogenic state

  1. Christian SM Helker  Is a corresponding author
  2. Jean Eberlein
  3. Kerstin Wilhelm
  4. Toshiya Sugino
  5. Julian Malchow
  6. Annika Schuermann
  7. Stefan Baumeister
  8. Hyouk-Bum Kwon
  9. Hans-Martin Maischein
  10. Michael Potente
  11. Wiebke Herzog
  12. Didier YR Stainier  Is a corresponding author
  1. Department of Developmental Genetics, Max Planck Institute for Heart and Lung Research, Germany
  2. Philipps-University Marburg, Faculty of Biology, Cell Signaling and Dynamics, Germany
  3. Angiogenesis and Metabolism Laboratory, Max Planck Institute for Heart and Lung Research, Germany
  4. University of Muenster, Germany
  5. DZHK (German Center for Cardiovascular Research), partner site Frankfurt Rhine-Main, Germany
  6. Max Planck Institute for Molecular Biomedicine, Germany
https://doi.org/10.7554/eLife.55589
Share this article
Cite this article
  1. Christian SM Helker
  2. Jean Eberlein
  3. Kerstin Wilhelm
  4. Toshiya Sugino
  5. Julian Malchow
  6. Annika Schuermann
  7. Stefan Baumeister
  8. Hyouk-Bum Kwon
  9. Hans-Martin Maischein
  10. Michael Potente
  11. Wiebke Herzog
  12. Didier YR Stainier
(2020)
Apelin signaling drives vascular endothelial cells toward a pro-angiogenic state
eLife 9:e55589.
https://doi.org/10.7554/eLife.55589
  2. Download BibTeX
  3. Download .RIS

Abstract

To form new blood vessels (angiogenesis), endothelial cells (ECs) must be activated and acquire highly migratory and proliferative phenotypes. However, the molecular mechanisms that govern these processes are incompletely understood. Here, we show that Apelin signaling functions to drive ECs into such an angiogenic state. Zebrafish lacking Apelin signaling exhibit defects in endothelial tip cell morphology and sprouting. Using transplantation experiments, we find that in mosaic vessels, wild-type ECs leave the dorsal aorta (DA) and form new vessels while neighboring ECs defective in Apelin signaling remain in the DA. Mechanistically, Apelin signaling enhances glycolytic activity in ECs at least in part by increasing levels of the growth-promoting transcription factor c-Myc. Moreover, APELIN expression is regulated by Notch signaling in human ECs, and its function is required for the hypersprouting phenotype in Delta-like 4 (Dll4) knockdown zebrafish embryos. These data provide new insights into fundamental principles of blood vessel formation and Apelin signaling, enabling a better understanding of vascular growth in health and disease.

Introduction

Endothelial cell sprouting is a fundamental process of physiological and pathological blood vessel growth. Attracted by growth factors such as vascular endothelial growth factor-A (VEGF-A) secreted from hypoxic tissues, endothelial cells (ECs) break out of the quiescent vessel wall to form new vessel branches (Ferrara et al., 2003; Koch and Claesson-Welsh, 2012). ECs with higher levels of VEGF-A signaling become invasive tip cells that lead new vascular sprouts, while neighboring ECs with lower VEGF-A signaling become trailing stalk cells (Gerhardt et al., 2003). This process is coordinated by Delta-like 4 (DLL4)/Notch signaling. Activation of Notch receptors by their ligand DLL4, expressed by tip cells, represses tip cell behavior in stalk cells (Hellström et al., 2007; Leslie et al., 2007; Siekmann and Lawson, 2007; Suchting et al., 2007). Loss of Notch signaling, on the other hand, causes excessive tip cell formation and vascular overgrowth (Hellström et al., 2007; Leslie et al., 2007; Siekmann and Lawson, 2007; Suchting et al., 2007).

Apelin (Apln) is a small secreted peptide, which was initially identified because of its inotropic activity (Szokodi et al., 2002). Apelin was subsequently described as a tip cell-enriched gene (del Toro et al., 2010). Apelin (Tatemoto et al., 1998), as well as the newly identified ligand Apela (Chng et al., 2013; Pauli et al., 2014), can both activate the Apelin receptor (Aplnr), a 7-transmembrane G-protein-coupled receptor (GPCR). Mouse and frog embryos lacking Apln or Aplnr function exhibit reduced vascular outgrowth, decreased EC proliferation, smaller vessel diameter as well as defects in the alignment of arteries and veins (Cox et al., 2006; Kälin et al., 2007; Kidoya et al., 2008; del Toro et al., 2010; Kidoya et al., 2010; Kidoya et al., 2015; Papangeli et al., 2016). In addition, Apelin signaling has been implicated in several cardiovascular diseases including pulmonary hypertension (Goetze et al., 2006; Alastalo et al., 2011; Chandra et al., 2011), atherosclerosis (Hashimoto et al., 2007; Chun et al., 2008; Kojima et al., 2010; Pitkin et al., 2010), myocardial infarction (Tempel et al., 2012; Wang et al., 2013; Zhang et al., 2016; Chen et al., 2017), and tumor angiogenesis (Kidoya et al., 2012; Zhao et al., 2018; Uribesalgo et al., 2019). However, the cellular mechanisms by which Apelin signaling functions within the vasculature remain elusive. Using zebrafish mutants combined with mosaic analyses, high-resolution time-lapse imaging, and metabolic studies, we find that Apelin signaling is required to boost endothelial metabolic activity during angiogenic sprouting. Furthermore, we show that Apelin signaling acts downstream of Notch signaling, where it is required for Notch-controlled angiogenesis.

Results

Apelin signaling is required for angiogenic sprouting

To examine the expression pattern of the apelin ligand and receptor genes during angiogenic sprouting in zebrafish embryos, we first performed whole-mount in situ hybridization during intersegmental vessel (ISV) formation. We detected clear alpn, but no apela, expression within the sprouting ISVs (Figure 1—figure supplement 1 arrowheads). For the receptor genes, we could only detect aplnrb expression in the ISVs (Figure 1—figure supplement 1 arrowheads).

To visualize apln and aplnrb expression at single cell resolution, we developed reporters using Bacterial artificial chromosome (BAC) recombineering (Figure 1—figure supplement 2). To this end, we replaced the ATG of an apln containing BAC with an EGFP cassette. Similarly, we replaced the stop codon of an aplnrb containing BAC with a tandem fluorescent timer (TagRFP-sfGFP) cassette leading to a fusion protein. We injected both modified BACs into one-cell stage zebrafish embryos to generate stable transgenic lines, Tg(apln:EGFP) and Tg(aplnrb:aplnrb-TagRFP-sfGFP) (hereafter referred to Tg(aplnrb:aplnrb-sfGFP)) (Figure 1—figure supplement 2). We first detected weak apln:EGFP expression in sprouting ISVs at 30 hpf (Figure 1A). At 54 hpf, all ECs within the dorsal longitudinal anastomotic vessel (DLAV) – a vessel formed by tip cells – were labeled (Figure 1A, arrowheads) while some stalk cells also exhibited weak apln:GFP expression (Figure 1A, arrows). Of note, aplnrb:Aplrnb-sfGFP expression at 26 hpf was visible in the entire ISV sprout (Figure 1B arrowheads), but it was absent from non-angiogenic ECs within the dorsal aorta (DA). At 54 hpf, aplnrb:Aplrnb-sfGFP expression was detected in all ECs that had sprouted out of the DA but also weakly in ECs within the DA (Figure 1B). These results suggest that apln is expressed in tip cells while aplnrb is expressed in all sprouting ECs.

Figure 1 with 3 supplements see all
Download asset Open asset
Apelin signaling promotes endothelial sprouting.

Visualization of apelin and apelin receptor b expression using transgenic reporter lines. Confocal projection images of the trunk region of zebrafish embryos. (A) TgBAC(apln:EGFP) expression is detectable in growing ISVs at 30 and 54 hpf. Arrowheads point to strong apln expression in tip cells, while arrows point to weak apln expression in stalk cells. (B) TgBAC(aplnrb:aplnrb-EGFP) expression is detectable in sprouting ECs (arrowheads) at 26 hpf and is clearly present in the ISVs and DLAV at 54 hpf. (C) Inactivation of Apelin ligand and receptor genes impairs angiogenesis. Confocal projection images of the blood vasculature in the trunk region of Tg(fli1a:EGFP) embryos. apln -/-; apela -/- as well as aplnra -/-; aplnrb -/- embryos exhibit a reduction in vascular sprouting at 28 and 54 hpf. Arrowheads point to stalled ISVs. Scale bars: A’, 10 µm; A’’, B’, C’, C’’, 20 µm; B’’, 15 µm.

To examine the function of Apelin signaling during sprouting angiogenesis in zebrafish, we used mutants for aplnra (Helker et al., 2015), aplnrb (Helker et al., 2015), apln (Helker et al., 2015) and apela (Chng et al., 2013). Homozygous aplnra mutant embryos exhibited no obvious defects during ISV formation (Figure 1—figure supplement 3). However, homozygous aplnrb mutant embryos exhibited reduced ISV length and failed to form the DLAV (Figure 1—figure supplement 3). This phenotype was more severe in embryos lacking both aplnra and aplnrb (Figure 1C, Figure 1—figure supplement 3), indicating partial compensation. We also analyzed apln and apela mutants. Homozygous apela mutant embryos displayed only a mild delay in ISV outgrowth (Figure 1—figure supplement 3), while homozygous apln mutant embryos exhibited defects in ISV outgrowth and failure to form the DLAV (Figure 1—figure supplement 3). Loss of both ligands increased the severity of the phenotype leading to ISV stalling at the horizontal myoseptum (Figure 1C, Figure 1—figure supplement 3). Consistent with studies in the mouse retina (del Toro et al., 2010), our studies identify apln expression as a marker of endothelial tip cells in zebrafish and show that Apelin signaling is required for angiogenic sprouting.

Apelin signaling regulates tip cell morphology

To investigate when the sprouting defects in Apelin signaling-deficient embryos first appear, we analyzed developmental time points when tip cells start to sprout out of the DA. However, no differences in sprout initiation or tip cell specification were observed in double homozygous receptor or ligand mutants (Figure 2—figure supplement 1, Figure 2A). Instead, we found that sprout elongation was slower in these mutant embryos, resulting in an overall reduction of sprout length (Figure 2A, Figure 2—video 1, Figure 2—video 2). Furthermore, while endothelial tip cells in wild-type embryos formed long filopodia which extended toward the dorsal side of the animal (Figure 2A I, II, Figure 2—video 1), aplnr mutant embryos (aplnra+/-; aplnrb -/- and aplnra -/-; aplnrb -/-) as well as aplnrb morpholino (MO) injected embryos (morphants) displayed a blunted tip cell morphology (Figure 2A III, 2A IV, 2B, C, Figure 2—video 2), a phenotype which did not recover over time.

Figure 2 with 5 supplements see all
Download asset Open asset
Apelin signaling regulates endothelial filopodia formation and endothelial cell numbers.

(A) Still images from confocal time-lapse movies of vascular development in wild-type and aplnra +/-; aplnrb -/- embryos. During sprouting, wild-type tip cells send out filopodia (arrowheads). aplnra +/-; aplnrb -/- embryos exhibit smaller sprouts and fail to form filopodia. (B) Confocal images of the blood vasculature in 24 hpf Tg(kdrl:HsHRAS-EGFP) embryos injected with Ctr MO and aplnrb MO. aplnrb morphant embryos exhibit smaller sprouts and fail to form filopodia (arrowheads). (C) aplnrb morphant embryos exhibit a reduction in the number of endothelial filopodia (Ctr MO, n = 10; aplnrb MO, n = 15). (D) Confocal images of the blood vasculature of 30 and 52 hpf Tg(fli1a:nEGFP) wild-type and aplnra +/-; aplnrb -/- embryos showing EC cell nuclei. (E) aplnra +/-; aplnrb -/- embryos exhibit reduced EC numbers in the ISVs (30 hpf: aplnr +/+, n = 3; aplnr +/-, n = 10; aplnr -/-, n = 8; 52 hpf: aplnr +/+, n = 5; aplnr +/-, n = 10; aplnr -/-, n = 9). n.s. not significant (two-tailed t-test). Scale bars: A, D, 20 µm; B, 40 µm; B, inset 10 µm.

Previously, we reported a role for Apelin signaling in establishing blood flow-induced EC polarity (Kwon et al., 2016). To determine whether the observed defects during sprouting were caused by defects in EC polarity, we analyzed the location of the Golgi apparatus during ISV formation in wild-type and mutant embryos. However, we could not detect obvious differences in EC polarity during angiogenic sprouting (Figure 2—figure supplement 2, arrowheads point to polarized ECs). Next, we asked whether Apelin signaling regulates the number of ECs, and so combined aplnr mutants with the Tg(fli1a:nEGFP) reporter line (Roman et al., 2002) to visualize EC nuclei. Compared to controls, aplnr mutants exhibited a reduction in ISV EC numbers of 1 cell at 30 hpf (4 instead of 5) and 2 cells at 52 hpf (5 instead of 7) (Figure 2D,E). We next assessed whether apln overexpression leads to ectopic sprouting. To this end, we generated an inducible transgenic line to overexpress apln under the control of the hsp70l promoter. However, global overexpression of apln did not lead to ectopic sprouting of blood vessels but led to mispatterned lymphatic vessels (Figure 2—figure supplement 3, arrows). Altogether, these data indicate that the angiogenic defects in Apelin signaling-deficient embryos are caused by filopodia defects and impaired cell migration. Apelin signaling also regulates the number of ECs within the ISV sprouts.

Apelin signaling drives the sprouting behavior of ECs

We hypothesized that aplnrb expression (Figure 1B) provides an advantage for ECs to sprout. To test this hypothesis, we generated chimeric embryos using wild-type and aplnr deficient embryos (Figure 3A). Upon transplantation of wild-type donor cells into wild-type hosts, 34,5% of the donor-derived ECs were present in the ISVs at 24 hpf (Figure 3B,C). In contrast, upon transplantation of wild-type donor cells into aplnr-deficient hosts, 80% of the donor-derived ECs were present in the ISVs at 24 hpf (Figure 3B,C). Together these data show that the apelin receptors function cell-autonomously in endothelial sprouting. The Apelin receptor has been shown to signal mainly through the G-protein Gαi (Habata et al., 1999). Therefore, we blocked Gαi function through the mosaic and vascular-specific overexpression of pertussis toxin (PTX). Our results show that ECs deficient for signaling though Gαi behave similarly to aplnr mutant ECs indicating that the Apelin receptor mediates its angiogenic effect through Gαi (Figure 3—figure supplement 1). Notably, wild-type donor-derived ECs in aplnr deficient embryos populated the entire dorsal part of the vasculature which is usually missing in these mutants, further confirming the cell-autonomous function of the Apelin receptors during angiogenesis (Figure 3—figure supplement 2). Together, these results indicate that apelin signaling primes ECs toward a sprouting state.

Figure 3 with 2 supplements see all
Download asset Open asset
Apelin signaling promotes the sprouting behavior of ECs.

(A) Experimental design: At the blastula stage, cells from Tg(kdrl:HsHRAS-mCherry) embryos were transplanted into host embryos obtained from Tg(fli1a:EGFP) aplnra +/-; aplnrb +/- incrosses. At 24 hpf, the mosaic embryos were imaged and the donor-derived ECs scored for their position. (B, C) 34,5% of wild-type donor-derived ECs in wild-type hosts were found within the ISVs. 80% of wild-type donor-derived ECs in aplnra +/-; aplnrb -/- hosts were found within the ISVs. Notably, wild-type ECs transplanted into aplnr- deficient embryos completely substituted for the lack of cells in the dorsal part of the vasculature at 54 hpf (Figure 3—figure supplement 2). Scale bars: B, 20 µm.

Apelin signaling functions downstream of Notch signaling

It has been previously reported that Notch-deficient ECs outcompete wild-type ECs during ISV sprouting (Siekmann and Lawson, 2007), an observation consistent with data in mouse (Jakobsson et al., 2010; Pitulescu et al., 2017). Since wild-type ECs similarly outcompete aplnr mutant ECs, we wanted to investigate potential links between Apelin and Notch signaling. Hence, we first blocked Notch signaling in TgBAC(apln:EGFP) embryos by injecting a dll4 MO. As previously reported (Leslie et al., 2007; Siekmann and Lawson, 2007), dll4 morphants exhibited a hypersprouting ISV phenotype (Figure 4A). Notably, we also observed a clear increase in apln:EGFP expression in the ectopic sprouts (Figure 4A). To test whether Apelin signaling is required as a downstream effector of Notch signaling during angiogenesis, we injected the dll4 MO into the offspring of apln and aplnrb heterozygous parents and compared the phenotype in homozygous mutant embryos versus their wild-type siblings. Strikingly, the hypersprouting phenotype of dll4 morphants was not present when Apln or Aplnrb function was lost (Figure 4B, Figure 4—figure supplement 1). To examine whether other hypersprouting phenotypes require Apelin signaling, we analyzed aplnrb, plexinD1 (plxnd1) double mutant embryos (Figure 4C,E). aplnrb mutant embryos exhibit reduced sprouting and plxnd1 mutant embryos exhibit ectopic sprouting in line with published data (Torres-Vázquez et al., 2004; Figure 4C,E). Loss of aplnrb function in the background of the plxnd1 mutant did not alter its hypersprouting phenotype (Figure 4C,E), suggesting that it is independent of Apelin function. Similar results were obtained when we analyzed ectopic venous sprouting in response to bmp2b overexpression (Wiley et al., 2011; Figure 4D,F). Together these data indicate that Apelin signaling is specifically required for Notch-modulated angiogenesis. To investigate whether apln expression is itself regulated by Notch signaling, we performed cell culture assays. We treated HUVECs with the Notch inhibitor DAPT and analyzed APLN expression by RT-qPCR. Consistent with the observations in zebrafish (Figure 4A), we observed an increase in APLN expression upon Notch inhibition (Figure 4G, Figure 4—source data 1). Next, we activated Notch signaling by stimulating HUVECs with the Notch ligand DLL4. Conversely to the Notch inhibition data, activating Notch signaling in HUVECs suppressed APLN expression (Figure 4H, Figure 4—source data 1). Together these data suggest that the increased sprouting in response to Notch inhibition is, in part, driven by the upregulation of apln.

Figure 4 with 1 supplement see all
Download asset Open asset
Apelin signaling functions downstream of Notch signaling in endothelial cells.

(A - D) Confocal projection images of the blood vasculature in the trunk region of Tg(flt1:tdTomato) (A) and Tg(fli1a:EGFP) (B–D) animals at 54 (B–D) and 72 (A) hpf. (A) Injection of a dll4 morpholino leads to an increase in TgBAC(apln:EGFP) expression. (B) Loss of Apelin function can block excessive endothelial sprouting in dll4 morphants. (C, E) Angiogenic response in aplnrb -/-, plxnd1 -/-, and aplnrb -/-; plxnd1 -/- embryos (arrowheads) (n = 95). (D, F) Angiogenic response to bmp2b overexpression in aplnr +/- and aplnr -/- embryos (arrowheads). (E) Genotype of embryos for aplnrb after sorting them according to phenotype. (G) RT-qPCR analysis of APELIN mRNA levels in HUVECs treated with DAPT for 24 hr. Blocking Notch signaling with DAPT induces APELIN expression. (H) RT-qPCR analysis of APELIN mRNA levels in HUVECs cultured on DLL4 to activate Notch signaling. Activating Notch signaling represses APELIN expression. Arrowheads point to ectopic sprouts. n.s. not significant (two-tailed t-test). Ct values can be found in Figure 4—source data 1. Scale bars: A, C, 20 µm; B, 15 µm; D, 30 µm.

Figure 4—source data 1

Ct values of RT-qPCR.

https://cdn.elifesciences.org/articles/55589/elife-55589-fig4-data1-v2.docx

Apelin signaling positively regulates EC metabolism

Because EC sprouting requires an increase in metabolic activity (Dobrina and Rossi, 1983; Krützfeldt et al., 1990; Mertens et al., 1990; De Bock et al., 2013a; Vandekeere et al., 2015) and Apelin signaling has been shown to control cell metabolism in other contexts (Dray et al., 2008; Sawane et al., 2013), we asked whether Apelin signaling promotes EC metabolism. Previous studies have demonstrated that ECs rely on glycolysis for sprouting (De Bock et al., 2013a; Vandekeere et al., 2015). Therefore, we measured the extracellular acidification rate (ECAR) as a surrogate parameter of glycolysis in Apelin signaling-deficient HUVECs (Figure 5A,B). Notably, we observed a marked reduction in glycolysis after knockdown of Apelin signaling (Figure 5A), whereas mitochondrial oxygen consumption appeared unchanged (Figure 5B). To gain insight into the underlying mechanisms, we analyzed key regulators of metabolism and found a reduction in c-MYC protein levels after depletion of Apelin signaling (Figure 5C). Furthermore, expression of PFKFB3, which encodes an enzyme that sustains high glycolytic rates, was also reduced in Apelin signaling-deficient HUVECs (Figure 5D). In order to analyze whether a reduction in EC metabolic activity causes the vascular phenotype observed in aplnrb mutants, we performed mosaic rescue experiments and overexpressed pfkfb3 in ECs. In agreement with our in vitro data, we found that overexpression of pfkfb3 in endothelial tip cells leads to a partial rescue of the vascular phenotype in aplnrb mutants (Figure 5E arrowheads, 5F, Figure 4—source data 1). Thus, Apelin signaling controls the expression of regulators of glucose metabolism as well as glycolytic activity in developing endothelial cells.

Figure 5 with 1 supplement see all
Download asset Open asset
Apelin signaling positively regulates EC metabolism.

(A - B) Extracellular acidification aate (ECAR) (A) and oxygen consumption rates (OCR) (B) in siScr and APLN+APLNR siRNA-treated HUVECs under basal conditions and in response to oligomycin, fluoro-carbonyl cyanide phenylhydrazone (FCCP) and antimycin A (AA)/rotenone. (A) Reduced basal and maximal glycolytic activity in APLN+APLNR siRNA-treated compared to siScr-treated HUVECs. (B) No significant difference in oxygen consumption in APLN+APLNR siRNA-treated compared to siScr-treated HUVECs. (C) Reduced c-MYC levels in APLN+APLNR siRNA-treated compared to siScr-treated HUVECs. (D) RT-qPCR analysis of PFKFB3 mRNA levels in APLN+APLNR siRNA-treated compared to siScr-treated HUVECs. (E) Confocal projection images of the blood vasculature in the trunk region of a 54 hpf Tg(kdrl:HsHRAS-mCherry) animal injected with an EGFP:fli1a:pfkfb3 plasmid. Arrowheads point to formed DLAV fragments while asterisks indicate missing DLAV fragments. (F) Quantification of the rescue of the DLAV fragment by mosaic pfkfb3 overexpression in aplnrb -/- embryos. n.s. not significant (two-tailed t-test). Scale bar: E, 50 µm.

Discussion

During the formation of the first embryonic blood vessels, angioblasts migrate to the midline where they coalesce to form the future DA and cardinal vein. We have previously reported that vasculogenesis relies on the function of the ligand Apela (Helker et al., 2015). Here, we show that angiogenesis depends mostly on the function of the ligand Apln. However, Apela can partially compensate for the loss of Apln. This stage-specific ligand usage is in agreement with previous studies showing that apela expression is reduced by the end of vasculogenesis when apln starts to be expressed (Chng et al., 2013; Pauli et al., 2014).

During angiogenesis in embryos lacking Apelin signaling, we observed a severe sprouting defect with a reduction in EC numbers and filopodia. As ECs proliferate, extend filopodia, and migrate during ISV formation, it is challenging to assign the cause of the sprouting defect to the EC proliferation or filopodia formation defects. However, Phng et al., 2013 reported that the inhibition of filopodia formation by Latrunculin B treatment reduces ISV sprout length, suggesting that the ISV sprouting defects in apln mutants is caused by the filopodia defects. However, one cannot exclude the possibility that defects in EC numbers are also contributing to the ISV sprouting defects.

While we observed a severe angiogenesis phenotype when Apelin signaling was impaired, global overexpression of apln did not lead to ectopic sprouting. However, these experiments were done in the presence of endogenous Apelin, and thus, it is possible that the endogenous Apelin gradient prevents ECs from ectopic sprouting. In addition, Apelin might need to be expressed from a discrete source, rather than globally, to elicit a sprouting response.

During sprouting angiogenesis, ECs within a sprout are highly heterogenous in their shape, gene expression and function, which led to the model of tip and stalk cells (Gerhardt et al., 2003). While differences in expression between tip and stalk cells have been reported for several genes (Tammela et al., 2008), (Hellström et al., 2007; Siekmann and Lawson, 2007; Leslie et al., 2007; Suchting et al., 2007; del Toro et al., 2010; Bussmann et al., 2011; Herbert et al., 2012), little is known about the molecular differences between sprouting and resting ECs (Schlereth et al., 2018). By analyzing novel reporter lines for apln and aplnrb expression, we observed high apln expression in tip cells while we could not observe any difference in aplnrb expression between tip and stalk cells (Figure 5—figure supplement 1). Interestingly, aplnrb is highly expressed in sprouting ECs in ISVs while being absent from non-angiogenic ECs in the DA (Figure 5—figure supplement 1). These observations are in line with a recent study showing that ECs during tumor angiogenesis can be labeled by a CreERT2 transgene in the Aplnr locus while quiescent blood vessels in the surrounding tissue are not labeled (Zhao et al., 2018).

At the molecular level, vascular sprouting and cell positioning within the sprout is tightly regulated by VEGF and Notch signaling (Hellström et al., 2007; Lobov et al., 2007; Siekmann and Lawson, 2007; Suchting et al., 2007; Jakobsson et al., 2010). In addition to these signaling pathways, we propose Apelin signaling as a molecular switch to drive ECs into a pro-angiogenic state. In line with the expression of aplnrb in sprouting but not quiescent ECs, we show that aplnrb function regulates the ability of ECs to sprout or stay quiescent. Similarly, Notch signaling regulates the behavior of ECs (Siekmann and Lawson, 2007): rbpj deficient ECs contribute to the ISVs while wild-type ECs stay within the DA (Siekmann and Lawson, 2007). Of note, we found that Notch signaling regulates the expression of apln in vitro as well as in vivo and that Apelin signaling is a key downstream effector of Notch signaling during angiogenesis (Figure 5—figure supplement 1). However, it is very unlikely that apln is a direct Notch target gene since activation of Notch signaling leads to a downregulation of APLN expression. Thus far, two downstream effectors of Notch signaling have been reported to control angiogenesis namely PTEN (Serra et al., 2015) and CXCR4, another GPCR (Hasan et al., 2017; Pitulescu et al., 2017). While PTEN has been shown to be required for Notch induced arrest in EC proliferation (Serra et al., 2015), CXCR4 mediates Notch-controlled EC migration (Hasan et al., 2017; Pitulescu et al., 2017). PTEN and Apelin both regulate AKT phosphorylation (Davies et al., 1998; Masri et al., 2004). Thus, one might speculate that AKT function is a common effector of PTEN and Apelin signaling in EC proliferation. Furthermore, we found that Apelin was required for EC migration in the absence of Notch signaling. Similarly, CXCR4 is required for EC migration in the absence of Notch signaling (Hasan et al., 2017; Pitulescu et al., 2017). CXCR4 and APLNR both signal through the G-protein Gαi (Moepps et al., 1997; Habata et al., 1999), and they might therefore have similar effects. Gpr124, another GPCR, has been reported to be required in tip cells during zebrafish angiogenesis (Vanhollebeke et al., 2015), similar to Aplnr. However, Gpr124 is required in tip cells only in the brain (Vanhollebeke et al., 2015), while Aplnr is required in tip cells in the ISVs, where it is most highly expressed.

Sprouting angiogenesis is controlled by genetically encoded signal transducers as well as by the metabolic state. However, how environmental signals modulate the metabolic activity of ECs is incompletely understood. Here, we show that Apelin signaling regulates the expression of PFKFB3 and c-MYC, two powerful drivers of EC metabolism (Wilhelm et al., 2016; De Bock et al., 2013b). Recently it has been shown that Apelin signaling promotes FOXO1 phosphorylation (Hwangbo et al., 2017), which negatively regulates its activity. Consistent with these findings, FOXO1 has been shown to suppress c-MYC expression (Wilhelm et al., 2016). Together these data raise the possibility that Apelin signals through FOXO1 to regulate c-MYC levels. Of note, genetic deletion of Pfkfb3 in mouse ECs leads to a reduction in their number as well as defects in filopodia formation and extension (De Bock et al., 2013b), phenocopying aplnr mutant embryos.

Taken together, our findings provide novel insights into a druggable pathway regulating angiogenesis and suggest that manipulating the angiogenic state of ECs by controlling Apln signaling might have therapeutic potential to control vascular growth in pathological settings.

Materials and methods

Key resources table
Reagent type
(species) or
resource		DesignationSource or
reference		IdentifiersAdditional
information
genetic reagent (D. rerio)Tg(fli1a:EGFP)y1Lawson and Weinstein, 2002ZFIN: y1
genetic reagent (D. rerio)Tg(fli1a:nEGFP)y7Roman et al., 2002,ZFIN: y7
genetic reagent (D. rerio)Tg(kdrl:HsHRAS-mCherry)s896Chi et al., 2008ZFIN: s896
genetic reagent (D. rerio)aplnramu296Helker et al., 2015ZFIN: mu296
genetic reagent (D. rerio)aplnrbmu281codes for another allele of aplnrb from Helker et al., 2015ZFIN: mu281
genetic reagent (D. rerio)aplnmu267Helker et al., 2015ZFIN: mu267
genetic reagent
(D. rerio)		Tg(hsp70:bmp2b)fr13Chocron et al., 2007ZFIN: fr13
genetic reagent (D. rerio)apelabr13Chng et al., 2013,ZFIN: br13
genetic reagent (D. rerio)Tg(fli:lifeact-GFP)mu240Hamm et al., 2016ZFIN: mu240
genetic reagent (D. rerio)Tg(fli1a:Hsa.B4GALT1-mCherry)bns9Kwon et al., 2016ZFIN: bns9
genetic reagent (D. rerio)Tg(hsp70:apln)mu269This manuscriptZFIN: mu269
genetic reagent (D. rerio)Tg(kdrl:HsHRAS-EGFP)mu280This manuscriptZFIN: mu280
genetic reagent (D. rerio)Tg(apln:EGFP)bns157This manuscriptZFIN: bns157
genetic reagent (D. rerio)Tg(aplnrb:aplnrb-TagRFP-
sfGFP)bns309		This manuscriptZFIN: bns309
antibodyanti-FOXO1 (rabbit monoclonal)Cell Signaling TechnologyCat#2880(1:1000)
antibodyanti-pThr24FOXO1/pThr32FOXO3a (rabbit monoclonal)Cell Signaling TechnologyCat#9464(1:1000)
antibodyanti-c-MYC (rabbit polyclonal)Cell Signaling TechnologyCat#9402(1:1000)
antibodyanti-Tubulin (rabbit polyclonal)Cell Signaling TechnologyCat#2148(1:1000)
otherTaqman probe APLNThermo Fisher ScientificHs00175572_m1
otherTaqman probe APLNRThermo Fisher ScientificHs00270873_s1
otherTaqman probe PFKFB3Thermo Fisher ScientificHs00270873_s1
otherTaqman probe ACTBThermo Fisher ScientificHs01060665_g1
commercial assay or kitIn-Fusion HD Cloning PlusTakara BioCat# 638910
transfected construct (human)APLNDharmaconCat# L-017023-01-000550 nM
transfected construct (human)APLNRDharmaconCat# L-005430-00-000550 nM
transfected construct (human)ON-TARGETplus Non-targeting PoolDharmaconCat# D-001810-10-0550 nM
commercial assay or kitmMessage mMachine SP6 Transcription KitThermo Fisher ScientificCat# AM1340
commercial assay or kitDIG RNA labelling kitRocheCat# 11277073910
commercial assay or kitSuperScript III First-Strand Synthesis SystemThermo Fisher ScientificCat#18080051
commercial assay or kitRNA Clean and Concentrator KitZymo ResearchCat# R1013
software, algorithmZEN Blue 2012Zeiss, Germany
software, algorithmZEN Black 2012Zeiss, Germany
software, algorithmImaris - Version 8.4.0Bitplane, UK
software, algorithmGraphPad Prism 6GraphPad Software, USA

Zebrafish husbandry and strains

Request a detailed protocol

All zebrafish housing and husbandry were performed under standard conditions in accordance with institutional (Max Planck Society) and national ethical and animal welfare guidelines approved by the ethics committee for animal experiments at the Regierungspräsidium Darmstadt, Germany, as well as the FELASA guidelines (Aleström et al., 2020). Embryos were staged by hours post fertilization (hpf) at 28.5°C (Kimmel et al., 1995). The following lines were used: Tg(fli1a:EGFP)y1 (Lawson and Weinstein, 2002), Tg(fli1a:nEGFP)y7 (Roman et al., 2002), Tg(kdrl:HsHRAS-mCherry)s896 (Chi et al., 2008), aplnramu296 (Helker et al., 2015), the aplnrbmu281 allele was generated using the same CRISPR as in Helker et al., 2015 and contains a 4 bp insertion 137 bp downstream of the ATG leading to a premature stop codon 196 bp downstream of the ATG, aplnmu267 (Helker et al., 2015), Tg(hsp70:bmp2b)fr13 (Chocron et al., 2007), apelabr13 (Chng et al., 2013), Tg(fli1a:LIFEACT-GFP)mu240 (Hamm et al., 2016), Tg(fli1a:Hsa.B4GALT1-mCherry)bns9 (Kwon et al., 2016), Tg(hsp70:apln)mu269 (this study), Tg(kdrl:HsHRAS-EGFP)mu280 (this study), Tg(apln:EGFP)bns157 (this study) and Tg(aplnrb:aplnrb-TagRFP-sfGFP)bns309 (this study).

Generation of the TgBAC(apln:EGFP)bns157, TgBAC(aplnrb:aplnrb-TagRFP-sfGFP)bns309, Tg(kdrl:HsHRAS-EGFP)mu280, and Tg(hsp70l:apln)mu269lines

Request a detailed protocol

To generate the apln and aplnrb bacterial artificial chromosome (BAC) constructs, we used the BAC clones RP71-2G21 containing the apln locus and CH211-102K containing the aplnrb locus. All recombineering steps were performed as described in Bussmann and Schulte-Merker, 2011 with the modifications as described in Helker et al., 2019. The following homology arms were used to generate the targeting PCR products of the EGFP_Kan, and TagRFP-sfGFP_Kan cassettes:apln-HA1: 5’-ccactacagtatatcagctagcgactggcagggaaacggaggggagagcaaccatggtgagcaagggcgaggag-3’ and apln-HA2: 5’-cacagcagagaaaccaccagcacaatcaccagcgtcaagatcttcacattttccagaagtagtgaggag-3’;aplnrb-HA1:5’-gctccctttcttcacagaagaccgaggcccagtcgctggctacgaaggtgcttggacctggactcggatc-‘3 and aplnrb-HA2: 5’-taattgctgacttgttaccccaattctgcgtcacccttccgttctcctcctgaccatgattacgccaagc-‘3.

To generate the Tg(kdrl:HsHRAS-EGFP) and Tg(hsp70:apln) lines, the gateway recombination system (Invitrogen) using entry vectors and the pTolDest destination vector (Villefranc et al., 2007) was used. The apln coding sequence was amplified from cDNA. 100 pg DNA of the plasmids and 50 pg of tol2 mRNA were injected into one-cell stage zebrafish embryos for stable germline transmission.

Morpholino injections

Request a detailed protocol

Morpholinos were obtained from Gene Tools, resuspended in distilled H2O and around 2 nl was injected into 1 cell stage embryos. The following morpholinos were used: aplnrb MO (Helker et al., 2015) at 0.5 ng/embryo, dll4 MO (Hogan et al., 2009) at 3 ng/embryo. An equal amount of the standard control MO: 5’-CCTCTTACCTCAGTTACAATTTATA-3’ was used for each experiment.

Transplantation experiments

Request a detailed protocol

At the sphere stage, cells were removed from Tg(kdrl:HsHRAS-mCherry) donor embryos and transferred to Tg(fli1a:EGFP) aplnr mutant hosts using a glass capillary. Transplanted ECs were identified by transgenic mCherry expression.

Whole-mount in situ hybridization

Request a detailed protocol

Single in situ hybridizations were performed as described (Thisse and Thisse, 2008; Helker et al., 2013). The following probes were synthesized: apln (Helker et al., 2015), apela (Chng et al., 2013), aplnra (Helker et al., 2015), and aplnrb (Helker et al., 2015).

Confocal microscopy

Request a detailed protocol

Zebrafish larvae were mounted in 1% low melt agarose. Egg water and agarose were supplemented with 19.2 mg/l Tricaine. All fluorescent images were acquired using an upright Zeiss LSM 780, 800 or 880 or a Leica SP5 or SP8 confocal microscope. Maximum projection images were analyzed and generated using Imaris (Bitplane).

Quantification of mutant phenotypes

Request a detailed protocol

For every embryo, somites 5 to 15 were analyzed (normal: 10 fully developed ISVs and connected DLAV; mild: 10 ISVs fully developed but no DLAV; strong: 1 to 6 ISVs shortened; severe: 1 to 10 ISVs shortened).

Quantification of filopodia

Request a detailed protocol

Only filopodia with more than 10 µm in length were used for quantification.

ISVs were categorized as filopodia rich ISVs (more than six filopodia) or filopodia poor ISVs (less than six filopodia). A total of 49 ISVs were quantified for the Ctr MO and 103 ISVs for the aplnrb MO.

pfkfb3 rescue experiments

Request a detailed protocol

pfkfb3 was cloned downstream of a bidirectional fli1a promoter driving EGFP in one direction and pfkfb3 in the other direction. aplnrb mutant embryos were injected with 20 pg EGFP:fli1a:pfkfb3 DNA and 30 pg Tol2 mRNA to generate mosaic blood vessels. EGFP positive tip cells were analyzed to quantify the percentage of connected DLAV segments. Neighboring EGFP negative tip cells were used as controls.

Cell culture

Request a detailed protocol

Pooled human umbilical vein endothelial cells (HUVECs) were purchased from Lonza (#CC-2519) and cultured in endothelial basal medium (EBM, Lonza) supplemented with hydrocortisone (1 μg/ml), bovine brain extract (12 μg/ml), gentamicin (50 μg/ml), amphotericin B (50 ng/ml), epidermal growth factor (10 ng/ml), and 10% fetal bovine serum (FBS, Life Technologies). HUVECs were tested for mycoplasma and cultured until the fourth passage. Cells were maintained at 37°C in a humidified atmosphere with 5% CO2.

RNA interference

Request a detailed protocol

To silence APLN and APLNR gene expression, HUVECs were transfected with 50 nM APLN and APLNR ON-TARGET SMARTpool siRNA (Dharmacon). As a control, a non-targeting siRNA pool was used (Dharmacon). HUVECs were grown to 70% confluency and transfected with Lipofectamine RNAiMAX (Life Technologies) according to manufacturer's instructions.

Western blot analysis and antibodies

Request a detailed protocol

Western blot analyses were performed with precast gradient gels (Bio-Rad) using standard methods. Briefly, cells were lysed in RIPA buffer (Sigma; 150 mM NaCl, 1.0% IGEPAL CA-630, 0.5% sodium deoxycholate, 0.1% SDS, and 50 mM Tris, pH 8.0) supplemented with Complete Protease Inhibitor Cocktail (Roche) and 1 mM PMSF. Proteins were separated by SDS-PAGE (Tris-glycine gels with Tris/glycine/SDS buffer, Bio-Rad) and transferred onto nitrocellulose membranes using the Trans Turbo Blot system (Bio-Rad). Membranes were probed with specific primary antibodies and then with peroxidase-conjugated secondary antibodies. The following primary antibodies were used: FOXO1 (Cell Signaling Technology, #2880, 1:1000), pThr24FOXO1/pThr32FOXO3a (Cell Signaling Technology, #9464, 1:1000), c-MYC (Cell Signaling Technology, #9402, 1:1000), Tubulin (Cell Signaling Technology, #2148, 1:1000), Secondary antibodies are peroxidase-conjugated Goat IgGs (1:5000) purchased from Jackson Immuno Research Labs. The target proteins were visualized by chemiluminescence using an ECL detection kit (Clarity Western ECL Substrate, Bio-Rad) and a ChemiDoc MP Imaging System (Bio-Rad).

RT-qPCR

Request a detailed protocol

Total RNA from HUVECs was extracted using a RNeasy Mini Kit (Qiagen). Reverse transcription polymerase chain reaction (RT-PCR) was performed using a SuperScript III First-Strand Synthesis System (Invitrogen) according to manufacturer’s instructions. RT-qPCR was carried out to quantify gene expression levels on a CFX connect Realtime System (Bio-Rad) with the following Taqman probes: APLN Hs00175572_m1, APLNR Hs00270873_s1, PFKFB3 Hs00270873_s1. Each sample was normalized to the housekeeping probe ACTB Hs01060665_g1.

Metabolic assay

Request a detailed protocol

The metabolism of cells was assessed by the measurement of extracellular acidification (ECAR) and oxygen consumption rates (OCR) using a Seahorse XFe96 analyser (Agilent). Four hours before the measurement, 40.000 HUVECs per well were seeded in a fibronectin-coated XFe96 microplate. The measurement was done following manufacturer’s protocol. To monitor glycolysis, the glycolysis stress test kit was used. The following substances were sequentially injected after a baseline measurement: Glucose (10 mM), Oligomycin (3 µM) and 2-Deoxyglucose (2-DG; 100 mM). The oxygen consumption rate was assessed using the Mito stress test kit. After a baseline measurement, the following substances were sequentially injected: Oligomycin (3 µM), the mitochondrial uncoupler carbonyl cyanide-4-(trifluoromethoxy)phenyl-hydrazone (FCCP; 1 µM) as well as a mixture of antimycin A (1.5 µM) and rotenone (3 µM).

Statistics

Standard error of the mean and P-values from a two-tailed t-test were calculated using Prism.

Data availability

All data generated or analysed during this study are included in the manuscript and supporting files.

References

    1. Davies MA
    2. Lu Y
    3. Sano T
    4. Fang X
    5. Tang P
    6. LaPushin R
    7. Koul D
    8. Bookstein R
    9. Stokoe D
    10. Yung WK
    11. Mills GB
    12. Steck PA
    (1998)
    'Adenoviral transgene expression of MMAC/PTEN in human glioma cells inhibits Akt activation and induces anoikis'
    Cancer Research 58:5285–5290.
    1. Roman BL
    2. Pham VN
    3. Lawson ND
    4. Kulik M
    5. Childs S
    6. Lekven AC
    7. Garrity DM
    8. Moon RT
    9. Fishman MC
    10. Lechleider RJ
    11. Weinstein BM
    (2002)
    'Disruption of acvrl1 increases endothelial cell number in zebrafish cranial vessels'
    Development 129:3009–3019.

Article and author information

Author details

  1. Christian SM Helker

    1. Department of Developmental Genetics, Max Planck Institute for Heart and Lung Research, Bad Nauheim, Germany
    2. Philipps-University Marburg, Faculty of Biology, Cell Signaling and Dynamics, Marburg, Germany
    Contribution
    Conceptualization, Supervision, Funding acquisition, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    For correspondence
    christian.helker@biologie.uni-marburg.de
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-0427-5338

  2. Jean Eberlein

    1. Department of Developmental Genetics, Max Planck Institute for Heart and Lung Research, Bad Nauheim, Germany
    2. Philipps-University Marburg, Faculty of Biology, Cell Signaling and Dynamics, Marburg, Germany
    Contribution
    Investigation, Writing - review and editing
    Competing interests
    No competing interests declared

  3. Kerstin Wilhelm

    Angiogenesis and Metabolism Laboratory, Max Planck Institute for Heart and Lung Research, Bad Nauheim, Germany
    Contribution
    Investigation, Writing - review and editing, equal contribution with Toshiya Sugino
    Competing interests
    No competing interests declared

  4. Toshiya Sugino

    Angiogenesis and Metabolism Laboratory, Max Planck Institute for Heart and Lung Research, Bad Nauheim, Germany
    Contribution
    Investigation, Writing - review and editing, equal contribution with Kerstin Wilhelm
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-6330-7275

  5. Julian Malchow

    Philipps-University Marburg, Faculty of Biology, Cell Signaling and Dynamics, Marburg, Germany
    Contribution
    Investigation
    Competing interests
    No competing interests declared

  6. Annika Schuermann

    University of Muenster, Muenster, Germany
    Contribution
    Investigation
    Competing interests
    No competing interests declared

  7. Stefan Baumeister

    Philipps-University Marburg, Faculty of Biology, Cell Signaling and Dynamics, Marburg, Germany
    Contribution
    Investigation, Writing - review and editing
    Competing interests
    No competing interests declared

  8. Hyouk-Bum Kwon

    Department of Developmental Genetics, Max Planck Institute for Heart and Lung Research, Bad Nauheim, Germany
    Contribution
    Investigation, Writing - review and editing
    Competing interests
    No competing interests declared

  9. Hans-Martin Maischein

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

  10. Michael Potente

    1. Angiogenesis and Metabolism Laboratory, Max Planck Institute for Heart and Lung Research, Bad Nauheim, Germany
    2. DZHK (German Center for Cardiovascular Research), partner site Frankfurt Rhine-Main, Berlin, Germany
    Contribution
    Supervision, Funding acquisition, Writing - review and editing
    Competing interests
    No competing interests declared

  11. Wiebke Herzog

    1. University of Muenster, Muenster, Germany
    2. Max Planck Institute for Molecular Biomedicine, Muenster, Germany
    Contribution
    Supervision, Funding acquisition, Writing - review and editing
    Competing interests
    No competing interests declared

  12. Didier YR Stainier

    1. Department of Developmental Genetics, Max Planck Institute for Heart and Lung Research, Bad Nauheim, Germany
    2. DZHK (German Center for Cardiovascular Research), partner site Frankfurt Rhine-Main, Berlin, Germany
    Contribution
    Conceptualization, Supervision, Funding acquisition, Writing - original draft, Project administration, Writing - review and editing
    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

Deutsche Forschungsgemeinschaft (SFB 834)

  • Christian SM Helker
  • Didier YR Stainier

Deutsche Forschungsgemeinschaft (GRK2213)

  • Christian SM Helker
  • Jean Eberlein
  • Julian Malchow

Deutsche Forschungsgemeinschaft (HE4585/3-1)

  • Wiebke Herzog

H2020 European Research Council (EMERGE (773047))

  • Michael Potente

Deutsche Forschungsgemeinschaft (EXC 2026)

  • Michael Potente

H2020 European Research Council (AdG project: ZMOD 694455)

  • Didier YR Stainier

North Rhine-Westphalia (Return fellowship)

  • Wiebke Herzog

Max-Planck-Gesellschaft

  • Christian SM Helker
  • Kerstin Wilhelm
  • Toshiya Sugino
  • Hyouk-Bum Kwon
  • Hans-Martin Maischein

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

Acknowledgements

We would like to thank Radhan Ramadass and the Marburg Center of Advanced Light Microscopy, Marianne Ploch and all the fish facility staff for technical support; Teja Mullapudi, Giulia Boezio, Jialing Qi and Constanze Heinzen for suggestions and critical comments on the project. Research in the Herzog laboratory was supported by the North Rhine-Westphalia 'return fellowship' and the DFG (HE4585/3-1). Research in the Potente laboratory is supported by the Max Planck Society, the European Research Council (ERC) Consolidator Grant EMERGE (773047), the DFG (SFB 834), the Excellence Cluster Cardio-Pulmonary Institute (EXC 2026, Project ID: 390649896), the DZHK (German Center for Cardiovascular Research), and a grant by the Leducq Foundation. Research in the Stainier laboratory is supported in part by the Max Planck Society, the DFG (SFB 834/4), the EU (ERC AdG project: ZMOD 694455) and the Leducq Foundation, and in the Helker laboratory by the DFG (SFB 834/4) and GRK2213.

Ethics

Animal experimentation: Ethics Statement All zebrafish husbandry was performed under standard conditions, and all experiments were conducted in accordance with institutional (MPG) and national ethical and animal welfare guidelines (Proposal numbers: B2/1017, B2/1041, B2/1218, B2/1138). All procedures conform to the guidelines from Directive 2010/63/EU of the European Parliament on the protection of animals used for scientific purposes.

Version history

  1. Received: January 29, 2020
  2. Accepted: September 19, 2020
  3. Accepted Manuscript published: September 21, 2020 (version 1)
  4. Version of Record published: October 16, 2020 (version 2)

Copyright

© 2020, Helker 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

  • 5,395
    Page views
  • 657
    Downloads
  • 49
    Citations

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

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. Christian SM Helker
  2. Jean Eberlein
  3. Kerstin Wilhelm
  4. Toshiya Sugino
  5. Julian Malchow
  6. Annika Schuermann
  7. Stefan Baumeister
  8. Hyouk-Bum Kwon
  9. Hans-Martin Maischein
  10. Michael Potente
  11. Wiebke Herzog
  12. Didier YR Stainier
(2020)
Apelin signaling drives vascular endothelial cells toward a pro-angiogenic state
eLife 9:e55589.
https://doi.org/10.7554/eLife.55589

Share this article

https://doi.org/10.7554/eLife.55589

Categories and tags

Research organism

Further reading

    1. Developmental Biology

    Inhibition of the serine protease HtrA1 by SerpinE2 suggests an extracellular proteolytic pathway in the control of neural crest migration

    Edgar M Pera, Josefine Nilsson-De Moura ... Ivana Milas
    Research Article

    We previously showed that SerpinE2 and the serine protease HtrA1 modulate fibroblast growth factor (FGF) signaling in germ layer specification and head-to-tail development of Xenopus embryos. Here, we present an extracellular proteolytic mechanism involving this serpin-protease system in the developing neural crest (NC). Knockdown of SerpinE2 by injected antisense morpholino oligonucleotides did not affect the specification of NC progenitors but instead inhibited the migration of NC cells, causing defects in dorsal fin, melanocyte, and craniofacial cartilage formation. Similarly, overexpression of the HtrA1 protease impaired NC cell migration and the formation of NC-derived structures. The phenotype of SerpinE2 knockdown was overcome by concomitant downregulation of HtrA1, indicating that SerpinE2 stimulates NC migration by inhibiting endogenous HtrA1 activity. SerpinE2 binds to HtrA1, and the HtrA1 protease triggers degradation of the cell surface proteoglycan Syndecan-4 (Sdc4). Microinjection of Sdc4 mRNA partially rescued NC migration defects induced by both HtrA1 upregulation and SerpinE2 downregulation. These epistatic experiments suggest a proteolytic pathway by a double inhibition mechanism:

    SerpinE2 ┤HtrA1 protease ┤Syndecan-4 → NC cell migration.

    1. Developmental Biology
    2. Neuroscience

    Fetal influence on the human brain through the lifespan

    Kristine B Walhovd, Stine K Krogsrud ... Didac Vidal-Pineiro
    Research Article

    Human fetal development has been associated with brain health at later stages. It is unknown whether growth in utero, as indexed by birth weight (BW), relates consistently to lifespan brain characteristics and changes, and to what extent these influences are of a genetic or environmental nature. Here we show remarkably stable and lifelong positive associations between BW and cortical surface area and volume across and within developmental, aging and lifespan longitudinal samples (N = 5794, 4–82 y of age, w/386 monozygotic twins, followed for up to 8.3 y w/12,088 brain MRIs). In contrast, no consistent effect of BW on brain changes was observed. Partly environmental effects were indicated by analysis of twin BW discordance. In conclusion, the influence of prenatal growth on cortical topography is stable and reliable through the lifespan. This early-life factor appears to influence the brain by association of brain reserve, rather than brain maintenance. Thus, fetal influences appear omnipresent in the spacetime of the human brain throughout the human lifespan. Optimizing fetal growth may increase brain reserve for life, also in aging.

    1. Developmental Biology
    2. Immunology and Inflammation

    Fetal liver macrophages contribute to the hematopoietic stem cell niche by controlling granulopoiesis

    Amir Hossein Kayvanjoo, Iva Splichalova ... Elvira Mass
    Research Article Updated

    During embryogenesis, the fetal liver becomes the main hematopoietic organ, where stem and progenitor cells as well as immature and mature immune cells form an intricate cellular network. Hematopoietic stem cells (HSCs) reside in a specialized niche, which is essential for their proliferation and differentiation. However, the cellular and molecular determinants contributing to this fetal HSC niche remain largely unknown. Macrophages are the first differentiated hematopoietic cells found in the developing liver, where they are important for fetal erythropoiesis by promoting erythrocyte maturation and phagocytosing expelled nuclei. Yet, whether macrophages play a role in fetal hematopoiesis beyond serving as a niche for maturing erythroblasts remains elusive. Here, we investigate the heterogeneity of macrophage populations in the murine fetal liver to define their specific roles during hematopoiesis. Using a single-cell omics approach combined with spatial proteomics and genetic fate-mapping models, we found that fetal liver macrophages cluster into distinct yolk sac-derived subpopulations and that long-term HSCs are interacting preferentially with one of the macrophage subpopulations. Fetal livers lacking macrophages show a delay in erythropoiesis and have an increased number of granulocytes, which can be attributed to transcriptional reprogramming and altered differentiation potential of long-term HSCs. Together, our data provide a detailed map of fetal liver macrophage subpopulations and implicate macrophages as part of the fetal HSC niche.